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C5: Membrane Permeability - Biology

C5: Membrane Permeability - Biology


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Another indicator of membrane dynamics is the measured permeability coefficient of ions and molecules across the bilayer. It is small and can enter down deep into the head group region through sequential H-bonding which must assist its transfer across the membrane.

Figure: Permeability of liposome bilayers

Many different probes have been developed to study membrane structure and dynamics. Many of them are fluorescent. The diagram below, taken from the Molecular Probe's catalog, gives examples.

Fluorescent Membrane Probes


Passive and Active Transport

Most biologically important solutes require protein carriers to cross cell membranes, by a process of either passive or active transport. Active transport requires the cell to expend energy to move the materials, while passive transport can be done without using cellular energy [4]. To put it another way, active transport uses energy to move a solute "uphill" against its gradient, whereas in facilitated diffusion, a solute moves down its concentration gradient and no energy input is required.

Therefore, to summarize, transport of solutes across cell membranes by protein carriers can occur in one of two ways [2]:

  • Downhill movement of solutes from regions of higher to lower concentration level, with the assistance of the protein carrier to pass through the membrane. This process is called passive transport or facilitated diffusion, and does not require energy.
  • Uphill movement of solute against the concentration gradient driving force (from regions of lower to higher concentration). Based on the chemical driving force, this process is unfavorable and requires some form of chemical energy to occur (active transport).

The type of transport process, facilitated/active transport, a biological cell employs is strictly dependent on its specific needs and concentration level of chemical/ions. For example, red blood cells use facilitated diffusion to transfer glucose across membranes, whereas intestinal epithelial cells rely on active transport to take in glucose from the gut [2]. Facilitated diffusion is effective for red blood cells primarily because the glucose concentration in the blood is stable and higher than the cellular level. In contrast, active transport is needed for the gut since there are large fluctuations of glucose level as a result of eating.

Figure (PageIndex<2>) encapsulates different transfer mechanisms discussed so far. Please note that the concentration gradient driving force is assumed downward in this schematic diagram.

Figure (PageIndex<2>) - Passive and active transports [3]

Facilitated Diffusion

In principle, there are two types of facilitated diffusion carriers as follows:

  1. Water molecules or certain ions can be transported by channel proteins. By forming a protein-lined pathway across the membrane, proteins can appreciably speed up the transfer rate of such solutes. It should be noted though that each type of channel protein is very selective to a specific ion/chemical. For example, some channels allow only K + ions to pass whereas they act like a barrier to other ions. Moreover, many of these channels are gated. To simply explain the issue, consider that the pathways are closed and unavailable for transport unless specific signals are given. One of the most vital functions of gated channels is in regulating nerve conduction in animals [2].
  2. Organic molecules, like sugars and amino acids, can be transferred across the membrane via uniporters which carry molecules along the concentration gradient. Almost all tissues in any living being have a variety of uniporters for transfer of glucose and amino acids into their cells.

Active Transport

Active transporters make an endergonic reaction (Keq < 1) more exergonic (Keq > 1) by coupling the first reaction to a second highly exergonic reaction (e.g., ATP-hyrolysis) through common intermediates to change the direction of transport (e.g., Na export from low to high concentration) [3]. To be more precise, when a transfer is not electrochemically favorable, another source of energy (which can come from another reaction) is required to force the transfer. These can be accomplished by a direct result of ATP hydrolysis (ATP pump) or by coupling the movement of one substance with that of another (symport or antiport) [2]. Active transport may use energy to transport solutes into or out of the cell, but always in opposite direction of the electrochemical driving force.

As mentioned before, biomembranes separate the intracellular and extracellular environments that are different in many aspects such as concentration levels of ions and chemicals. For example, in human tissues, all cells have a higher concentration level of sodium ion outside the cell than inside, while the exact opposite condition is maintained for the potassium ion (Cinside > Coutside). Regarding charged solutes and ions, besides the concentration gradient, the electrical voltage can come to play too there is an electrical driving force for cations and anions to move along and opposite the electric field, respectively.

Like pushing an object uphill against the gravitational field, moving a molecule against its favorable electrochemical driving force requires energy. In this respect, biological cells have evolved active protein transporters that can transfer ions and charged molecules in an electrochemically unfavorable direction.

In theory, active transport can be explained by a simple fact: Standard Free Energy Changes are Additive. Consider two reactions:

This rule can show how an endergonic reaction (Keq < 1) can shift to the RHS (producing more product) by being coupled to another highly exergonic reaction (Keq >> 1) through a common intermediate [3]. To clarify the issue let us consider the active transport of the sodium ion as follows:

The ion transport equation can be written as

[ Delta G = RT ln dfrac + zFV]

which for Na + gives the Gibbs free energy of 2.98 kcal/mol, or equivalently, an equilibrium constant of 0.0065. In this equation, R is the universal gas constant (1.987 cal/(mol.K)), T is absolute temperature (K), F is the Faraday's constant (23060 cal/(volt.mol)) and z is the the valence (charge number) of the ion. Moreover, subscripts i and o denote the inside and outside of the cell.

As mentioned before, the required excess energy for active transport can come from ATP hydrolysis. Typical Gibbs free energy change for ATP hydrolysis is around -13 kcal/mol, making the total Gibbs free energy change of 2.98 - 13 = -10.02 kcal/mol. Therefore, the overall reaction is highly shifted to produce more product by being coupled with the strictly exothermic ATP hydrolysis reaction.

Osmosis: Water Permeability

Osmosis (transfer of water molecules through the bilayer) is a function of the relative concentration levels of solute molecules in intracellular and extracellular environments. Water molecules can readily pass through special protein channels. If the total concentration of all dissolved solute is unbalanced (Cinside

= Coutside), there would be a net water flow into or out of the biological cell [5]. The direction and magnitude of the water flow is strictly dependent on whether the cell&rsquos environment is isotonic, hypotonic, or hypertonic which are illustrative measures for the relative concentrations of solutes inside and outside the cell.

Isotonic Solutions (Cinside = Coutside)

In isotonic case, the total molar concentration of dissolved solutes is the same for the intracellular and extracellular environments. In this condition, the inward and outward flows of water molecules are exactly balanced (shown in Figure (PageIndex<4>)). As shown in Figure (PageIndex<4>), the net flow of water is zero and total number of water molecules (or equivalently water concentration, Cw) is remained constant on each side. A 0.9% solution of sodium hydroxide is a perfect example of isotonic solution to animal cells [2]. During experiments, like exposing membranes to different solutions, it is highly recommended to use an isotonic solution to prevent osmotic effects (e.g., swelling and shrinking of the cell) which can seriously damage the biological cells.

Figure (PageIndex<4>) - Transport of water molecules through protein channels in isotonic condition [2]

Hypotonic Solutions (Cinside > Coutside)

In a hypotonic condition, molar concentration of the total dissolved solutes is higher inside the cell than that in the extracellular environment. Obviously, low concentration of solutes in an aqueous solution can be interpreted as high concentration of water. Therefore, it is straightforward to see if Cinside > Coutside &rarr Cw, inside < Cw, outside, providing a driving force for a net inward water flow to the cell. Hence, when a cell is exposed to such hypotonic conditions, there is net water movement into the cell and passing time, the concentration of water molecules inside the cell would be increased. Because of this considerable accumulation of water molecules, cells will swell and may even burst if the excess accumulated water is not removed from the intracellular environment.

Hypertonic Solutions (Cinside < Coutside)

Cell behavior under hypertonic condition is exactly the opposite of what explained for the hypotonic case (Cinside < Coutside &rarr Cw, inside > Cw, outside). In this case, the water concentration is higher in cell's interior than in its outside, so there would be a net outward water flow from the cell. Therefore, passing time, the water concentration level will decrease inside the cell and cell will shrink. As an important consequence of the low water level, the ability of cell to function or divide would be gradually lost [2]. It is interesting that hypertonic solutions like concentrated syrups have been used since ancient times for food preservation. This can be explained through the fact that microbial cells that would cause spoilage are dehydrated in these very hypertonic environments and would be unable to function [2].


Results

Study overview

The aim of this study was to develop an accessible assay for assessing cell permeabilisation for in vitro analysis, such that it is applicable to eukaryotic or prokaryotic cells, and for multiple macromolecules. To achieve this, we hypothesised that: 1) permeabilisation could be defined in terms of a general macromolecule size feature, such as MW. 2) An intrinsic cellular factor could serve as an internalisation marker for molecules of different MW.

To test these hypotheses, we sought an easily accessible internalisation marker and found SAv to be an ideal candidate. As such, we used SAv to design an easily accessible assay that, following permeabilisation, only requires labelling of cells with the SAv-conjugate of choice. Labelled cells can be analysed by flow cytometry (as performed here) or by another suitable SAv detection method available to researchers. The workflow for this assay is illustrated in sFigure 1a. In this setting, the SAv-conjugate will only bind to cell intrinsic biotin if the membrane/envelope permeabilisation has been successful. Thus, the internalisation of molecules with similar MWs should be feasible with the same permeabilisation method. This approach was tested here in bacterial (E. coli) and mammalian (4 T1) cells, with 4 different permeabilisation agents and 2 different SAv conjugates. In addition, the assay was validated with a functional experiment assessing the internalisation and activity of a nuclease with dimensions similar to those of the SAv conjugate tested, as described below.

SAv allows the assessment of membrane permeabilisation

Streptavidin-Cy5 served as the cell internalisation marker for Benzonase, a dimeric nuclease, with a MW of 60 KDa [23]. Each monomer corresponds to the dimensions of DNase I – a compact monomer with a MW of

30 KDa and dimensions of 4.6 × 4 × 3.5 nm [24]. Benzonase was the most cost effective enzyme of 6 DNases examined (sFigure 2b), it exhibited the highest levels of activity per quantity of enzyme used. The permeabilisation capability of FF and 4 non-ionic detergents for 60 KDa molecules was assessed in 4 T1 and E. coli cells following the workflow in sFigure 1a. As shown in Fig. 1a, impermeabilised FF 4 T1 cells showed significantly less SAv-Cy5 fluorescence than those exposed to detergents, while only E. coli cells (Fig. 1b) treated with Triton-X were permeabilised, as evidenced by a 363X increase in fluorescence (p < 0.001). This indicates that fixation does not permeabilise cells to large molecules. Among the detergents tested, Quillaja bark Saponin (Qb-Saponin) (displayed the highest membrane selectivity for Sav in 4 T1 cells, with a 186X (p < 0.001) increase in fluorescence, however no significant fluorescence change for E. coli was detected (p > 0.05). This held true for E. coli cells exposed to higher Qb-Saponin concentrations (sFigure 2a). When a larger molecule was examined in 4 T1 cells - 360 KDa Streptavidin Phycoerythrin (SAv-PE) – internalisation was much lower (2–25%) than that observed for SAv-Cy5, as expected, although patterns of detergent efficacy varied, with internalisation only detectable for Digitonin (10.8%) and Qb-Saponin (25%) (sFigure 3).

Membrane permeabilisation. Cell permeabilisation is measured by the internalisation of SAv-Cy5 for (a) 4 T1 cells and (b) E. coli. (Left) Histograms showing Cy5+ maximum fluorescence intensity (n = ( overline> ) 6). (Right) Box plot showing median fluorescence intensity. Deviation (%) from impermeabilised shown above each box in blue/red and p-values are shown in black. In all cases n = 6

Validation of the permeabilisation strategy by nuclease activity

Nuclease activity in permeabilised cells was tested by measuring the fluorescence emitted by a cell permeable, double-stranded DNA intercalating dye (CytoPhase Violet), after treatment with a permeabilisation (P+) agent and Benzonase (sFigure 1b). Following membrane permeabilisation, the nuclease can passively diffuse through the cytoplasm and pores in the nuclear membrane [25]. A reduction in CytoPhase signal is indicative of a reduction in DNA content, and thus higher nuclease activity. Results presented in Fig. 2 reflect those in Fig. 1, with the most significant CytoPhase signal reduction (30.8%, p < 0.001) observed in 4 T1 cells permeabilised with Qb-Saponin. Conversely, Qb-saponin treatment did not lead to any significant decrease in fluorescence for E. coli (4.5% decrease, p > 0.05), while treatment with Triton-X (P+ DNAse+ control), showed the greatest decrease 43.7% (p < 0.001). It was also noticeable that harvesting or pre-treatments did not significantly affect the integrity of the E. coli cells envelope, as impermeabilised cells exposed to Benzonase did not show a significant decrease in CytoPhase signal. These results were verified by qPCR, whereby a mixed FF cell population, containing 1 × 10 7 E. coli and 1 × 10 6 4 T1 cells, were exposed to the Host DNA depletion (HD) strategy, after which cells were harvested and DNA purified. Eluted DNA was analysed by qPCR. As seen in Fig. 3(i), for 4 T1 cells, the quantity of genes (normalised to genome copies) retrieved after HD were reduced by 10-fold (p < 0.01), which suggests that approximately 90% of the cells were permeabilised and their DNA content digested. Due to the reduced interference of mammalian DNA, HD treatment allowed for a higher (truer) representation of bacterial DNA, which exhibited a 3X (p < 0.01) increase in the number of genomes recovered (Fig. 3(ii)). Altogether, these results validate the permeabilisation assessment strategy and confirm that Qb-Saponin shows the best cell selective permeabilisation capacity.

DNA depletion. DNA depletion is measured here by a reduction in fluorescence of the double-stranded DNA intercalating dye CytoPhase, measured for (a) 4 T1 cells and (b) E. coli. (Left) Histograms showing the maximum fluorescence intensity for CytoPhase+ cells. (Right) Box plot showing median fluorescence intensity. Deviation (%) from impermeabilised DNase- shown above each box in blue/red and p-values are shown in black. In all cases n = 6

Quantifying DNA Depletion. DNA depletion measured by a reduction in the qPCR recovery of genomes of (i) 1 × 10 5 4 T1 cells and (ii) 1 × 10 6 E. coli from a mixed cell suspension treated or not with Qb-Saponin and Benzonase. Deviation (%) from impermeabilised DNase- shown above each box in blue/red and p-values are shown in black. In all cases n = 6


Bacterial killing by complement requires membrane attack complex formation via surface-bound C5 convertases

The immune system kills bacteria by the formation of lytic membrane attack complexes (MACs), triggered when complement enzymes cleave C5. At present, it is not understood how the MAC perturbs the composite cell envelope of Gram-negative bacteria. Here, we show that the role of C5 convertase enzymes in MAC assembly extends beyond the cleavage of C5 into the MAC precursor C5b. Although purified MAC complexes generated from preassembled C5b6 perforate artificial lipid membranes and mammalian cells, these components lack bactericidal activity. In order to permeabilize both the bacterial outer and inner membrane and thus kill a bacterium, MACs need to be assembled locally by the C5 convertase enzymes. Our data indicate that C5b6 rapidly loses the capacity to form bactericidal pores therefore, bacterial killing requires both in situ conversion of C5 and immediate insertion of C5b67 into the membrane. Using flow cytometry and atomic force microscopy, we show that local assembly of C5b6 at the bacterial surface is required for the efficient insertion of MAC pores into bacterial membranes. These studies provide basic molecular insights into MAC assembly and bacterial killing by the immune system.

Keywords: Gram‐negative bacteria atomic force microscopy complement convertase membrane attack complex.

© 2019 The Authors. Published under the terms of the CC BY 4.0 license.

Conflict of interest statement

The authors declare that they have no conflict of interest.

Figures

Figure 1. MAC in serum perturbs both…

Figure 1. MAC in serum perturbs both the outer and inner membrane of Gram‐negative bacteria

Schematic representation of engineered perimCherry/cytoGFP E. coli cells that express mCherry in the periplasmic space (between the outer and inner membrane) and GFP in the cytosol.

Structured illumination microscopy image of perimCherry/cytoGFP E. coli confirming localization of mCherry (red) in the periplasm and GFP (green) in the cytosol. Scale bar = 3 μm.

Outer membrane damage (mCherry intensity) and inner membrane damage (% Sytox positive) of perimCherry/cytoGFP E. coli bacteria exposed to (different concentrations of) human serum. Inner membrane damage correlates with killing (samples where bacteria are killed are indicated with gray shadings and a cross, see CFU data in Fig EV1B).

(D) Serum‐induced inner membrane damage (% Sytox positive) and (E) killing (CFU/ml) of different Gram‐negative strains depends on MAC components C5 and C8, but not on lysozyme (10% serum). Dotted line represents the detection limit of the assay.

Figure EV1. Serum‐induced inner membrane damage is…

Figure EV1. Serum‐induced inner membrane damage is essential for bacterial killing

Representative flow cytometry plots…

Representative flow cytometry plots of perimCherry/cytoGFP E. coli after 30 min of exposure to buffer or 10% human serum.

Bacterial viability (via colony enumeration on agar plates) of perimCherry/cytoGFP E. coli exposed to a concentration range of serum (samples identical to Fig 1C).

Successful depletion of serum from lysozyme (lysozyme‐specific ELISA black line), but sustained complement activity (CH50 red).

Figure 2. Purified MAC components lack the…

Figure 2. Purified MAC components lack the bactericidal activity of serum

Purified MAC (denoted as C5b6MAC) can be formed by mixing preassembled C5b6 complexes with C7, C8, and C9.

Lysis of human erythrocytes after exposure to a concentration range of preassembled C5b6 in the presence of 100 nM C7. After washing, erythrocytes were exposed to 20 nM C8 and 100 nM C9 for 30 min after which the OD405 nm of the supernatant was measured.

Bacterial viability of three Gram‐negative strains after exposure to buffer, 10% human serum or C5b6MAC. Buffer and serum conditions are the same as Fig 1E.

Permeabilization of the outer, but not inner membrane of perimCherry/cytoGFP E. coli cells exposed to C5b6MAC (different concentrations of C5b6 with fixed concentrations of C7‐C9).

Inner membrane damage of three Gram‐negative strains exposed to buffer, 10% serum or C5b6MAC. Buffer and serum conditions are the same as Fig 1D.

Figure EV2. MAC assembled from purified C5b6…

Figure EV2. MAC assembled from purified C5b6 lacks bactericidal activity

Lysis of liposomes after exposure…

Lysis of liposomes after exposure to preassembled C5b6 with or without C7, C8, and C9. Calcein release from liposomes was determined by measuring absorbance at OD340 nm 0.5% Triton X‐100 was used as a positive control.

Percentage lysis of rabbit erythrocytes exposed to buffer or C5b6MAC, compared to Milli‐Q (MQ) water as control (set at 100% lysis).

Killing of E. coli MG1655 after exposure to preassembled C5b6 with or without C7, C8, and C9 (at concentrations similar to (B) or at concentrations exceeding those of 100% serum (highlighted by an arrow) ± 100 nM C5b6, 600 nM C7, 350 nM C8, 900 nM C9).

Figure 3. Reconstituting bactericidal MAC assembly via…

Figure 3. Reconstituting bactericidal MAC assembly via surface‐bound C5 convertases

Schematic overview for Conv‐MAC formation.…

Schematic overview for Conv‐MAC formation. Bacteria were labeled with C5 convertases by pre‐incubation with C5‐deficient serum (Fig EV3). Following a washing step (@), convertase‐labeled bacteria were incubated with uncleaved C5, C6, C7, C8, and C9 (termed “Conv‐MAC”).

Bacterial viability of convertase‐labeled bacterial strains exposed to buffer (Conv) or C5‐C9 (Conv‐MAC).

Bacterial viability of convertase‐labeled E. coli MG1655 exposed to a concentration range of C5 in the presence of 100 nM C6, 100 nM C7, 20 nM C8, and 100 nM C9. “Ctrl” indicates bacteria that are pretreated with heat‐inactivated ΔC5 serum. Dotted line represents the detection limit of the assay.

Bacterial viability of convertase‐labeled E. coli MG1655 exposed to C5‐C9 or conditions lacking one MAC component. As an extra control, convertase formation was blocked during ΔC5 serum incubation by adding 5 μM compstatin.

Bacterial viability of E. coli MG1655 exposed to FB depleted serum in the presence of 20 μg/ml OmCI (to deposit C4b and C3b without Bb). After washing, bacteria were exposed to C5‐C9 in the presence or absence of C1 and C2 (to generate classical pathway C5 convertases, C4b2aC3b).

Figure EV3. Successful labeling of the bacterial…

Figure EV3. Successful labeling of the bacterial surface with C5 convertases

Schematic overview of complement activation and MAC formation on the bacterial membrane. Different recognition pathways (classical, lectin, and alternative) generate C3 convertases (C4b2a in the classical/lectin pathway, C3bBb in the alternative pathway) on the target cell surface that cleave the major complement protein C3 into C3b. C3b covalently attaches to the cell surface via a reactive thioester. At high C3b densities, C3 convertases associate with deposited C3b to form C5 convertases (C4b2aC3b in the classical/lectin pathway, C3bBbC3b in the alternative pathway). The C5 convertase then catalyzes conversion of C5 into C5a and C5b. C5b triggers MAC formation by sequential binding to C6, C7, C8, and multiple copies of C9.

Incubation of E. coli MG1655 with a concentration range of C5‐depleted serum (ΔC5 serum) results in surface labeling with alternative pathway convertases (C3bBbC3b, evidence by flow cytometry analysis of surface‐bound C3b and Bb).

Successful labeling of E. coli with C5 convertases. (C) E. coli MG1655 was pre‐incubated with 10% ΔC5 serum (labeled as: C5 convertase), heat‐inactivated ΔC5 serum (Ctrl), or ΔC5 serum supplemented with 5 μM compstatin (Ctrl). After washing, C3b deposition was measured by flow cytometry. (D) Bacteria were labeled as described in (C) and (after washing) incubated with a concentration range of C5. Release of C5a into supernatants was measured by a calcium flux‐based assay (Bestebroer et al, 2010).

Figure 4. MAC assembly via surface‐bound C5…

Figure 4. MAC assembly via surface‐bound C5 convertases leads to inner membrane damage

Outer membrane damage (mCherry intensity) and inner membrane damage (% Sytox positive) of convertase‐labeled perimCherry/cytoGFP E. coli cells incubated with a concentration range of C5 and fixed concentrations of C6‐C9.

Inner membrane damage of perimCherry/cytoGFP E. coli exposed to a concentration range of ΔC5 serum and, after washing, to C5‐C9. As controls, bacteria were incubated with heat‐inactivated ΔC5 serum or 5 μM compstatin was added to the ΔC5 serum to block C3b deposition.

Inner membrane damage of three different convertase‐labeled bacteria exposed to buffer (Conv) or C5‐C9 (Conv‐MAC).

Confocal microscopy images of convertase‐labeled perimCherry/cytoGFP E. coli exposed to buffer (Conv) or C5‐C9 (Conv‐MAC). Unlabeled bacteria exposed to 1% serum served as control. Green = GFP, red = To‐pro‐3 DNA dye. Scale bars = 3 μm.

Figure 5. Local assembly of C5b6 by…

Figure 5. Local assembly of C5b6 by surface‐bound C5 convertases is required for killing

Schematic overview of MAC assembly on convertase‐labeled bacteria by C5b6 that is locally generated by incubation with C5 and C6 (top) or by preassembled C5b6 (bottom).

Bacterial viability of convertase‐labeled E. coli MG1655 exposed to Buffer (Conv), preassembled C5b6 (Conv + C5b6MAC) or a mixture of C5 and C6 (Conv‐MAC), in the presence of C7, C8, and C9. Dotted line represents the detection limit of the assay.

(C) Inner membrane damage (% Sytox positive) and (D) outer membrane damage (mCherry) of convertase‐labeled perimCherry/cytoGFP E. coli exposed to a concentration range of preassembled C5b6 or a mixture of C5 and C6, in the presence of 100 nM C7. After washing, bacteria were exposed to 20 nM C8 and 100 nM C9.

Figure EV4. Local assembly of C5b6 by…

Figure EV4. Local assembly of C5b6 by surface‐bound C5 convertases is not essential to lyse…

Figure 6. C5b6 rapidly loses the capacity…

Figure 6. C5b6 rapidly loses the capacity to form bactericidal pores

Step‐wise assembly of MAC on convertase‐labeled bacteria. Convertase‐labeled bacteria were incubated with C5/C6 or C5/C6/C7 for 15 min, and subsequently washed (@) or treated with 10 μg/ml Eculizumab (Ecu). Then, the remaining MAC components (C7‐9 for C5/C6 or C8‐9 for C5/C6/C7, respectively) were added to the incubation mixture. In the control conditions (Conv‐MAC), the remaining MAC components were added to the incubation mixture without washing or adding an inhibitor. (A) Outer membrane damage (mCherry) and (B) inner membrane damage (% Sytox positive) were determined.

Figure 7. Inner membrane damage is driven…

Figure 7. Inner membrane damage is driven by MAC assembly at the outer membrane

Outer and inner membrane damage of convertase‐labeled bacteria exposed to different combinations of MAC components. “@” indicates a washing step.

Outer and inner membrane damage of convertase‐labeled bacteria exposed to C5‐C8 and after washing, to a concentration range of C9.

Figure 8. Local formation of C5b6 is…

Figure 8. Local formation of C5b6 is required for efficient insertion of MAC pores into…

Convertase‐labeled bacteria were exposed to a concentration range of either preassembled C5b6 (C5b6MAC) or a mixture of C5 and C6 (Conv‐MAC), in the presence of 100 nM C7. After washing, 20 nM C8 and 100 nM C9‐Cy3 were added. Controls at 0 nM C5b6 or C5‐C6 confirm that the detected C9‐Cy3 deposition is specifically related to MAC formation (solid lines). Proper insertion of pores was assessed by a previously described shaving method with trypsin (Moskovich & Fishelson, 2007). Bacteria were first incubated with MAC components for 30 min and subsequently treated with 20 μg/ml trypsin for 15 min at 37°C (dotted lines).

Convertase‐labeled perimCherry/cytoGFP bacteria (Green) exposed to C5b6MAC or Conv‐MAC. Conditions were similar to those in (A) however, C9‐Cy5 was used to detect MAC pores (Red). 100 nM of C5 and C6 or C5b6 was used in combination with 100 nM C7, 20 nM C8, and 100 nM C9‐Cy5. Conv + C5b6MAC and Conv‐MAC images were taken in separate experiments in which laser settings were adjusted to the staining intensity of C9‐Cy5 to properly visualize pore distribution. Scale bars = 3 μm.

Atomic force microscopy analysis of E. coli BL21 and MG1655 immobilized using the Poly‐L‐Lysine protocol. (C) Entire bacteria and high‐resolution comparisons of untreated and convertase‐labeled E. coli BL21 exposed to C5‐C9 (Conv‐MAC) for 10 min. Scale bars: 800 nm (left) and 30 nm (right). Height scales: 1 μm (left), 8 nm (top right), 22 nm (bottom right). Width of magnification boxes: 42 nm, height scales: 8 nm (top) and 13 nm (bottom). Arrows highlight E. coli porin structures an asterisk highlights the C5b‐7 stalk. Height profiles (bottom) are shown for the white dashed lines in the images. (D) Atomic force microscopy (height and phase images) of convertase‐labeled E. coli MG1655 exposed to a mixture of C5 and C6 (Conv‐MAC) or preassembled C5b6 (Conv + C5b6MAC), in the presence of C7, C8, C9, FB, and FD. Images were generated in the same experiment. Scale bars: 50 nm. Height scales: 15 nm. This figure and three other replicates are included in Fig EV5B and C.

Figure EV5. Local assembly of C5b6 is…

Figure EV5. Local assembly of C5b6 is required for stable insertion of MAC pores and…

Atomic force microscopy height (left) and phase (right) images of E. coli MG1655 incubated with buffer (untreated), convertases, and convertases plus either C5‐C8 (Conv + C5‐C8) or C5‐C9 (Conv‐MAC). Scale bars: 50 nm. Height scales: 5 nm (untreated), 9 nm (convertase), and 6 nm (conv + C5‐C8/MAC).

Atomic force microscopy height (B) and phase (C) images of convertase‐labeled E. coli MG1655 exposed to either C5 and C6 or preassembled C5b6 (Conv‐MAC versus Conv + C5b6MAC) in the presence of C7‐C9, FB, and FD. Data shown correspond to four separate experiments in each experiment, Conv‐MAC and Conv + C5b6MAC were compared directly. The upper images of (B and C) are also presented in Fig 8D. Scale bars: 50 nm. Height scales: 15 nm.

(D) Outer membrane damage (mCherry) and (E) inner membrane damage (% Sytox positive) of non‐opsonized or convertase‐labeled bacteria incubated with 10 nM of C5 and C6 or C5b6 in the presence of 10 nM C7. After washing, a concentration range of C8 and 100 nM C9 was added. Data represent mean ± SD of 3 independent experiments.

Figure 9. Structural model for C5b6 assembly…

Figure 9. Structural model for C5b6 assembly by C5 convertases

Hypothetical model for C5 cleavage…

Hypothetical model for C5 cleavage by the alternative pathway C5 convertase. The AP C5 convertase is a multimeric complex between a dimeric C3 convertase enzyme (comprised of surface‐bound non‐catalytic C3b in complex with protease Bb), together with additional surface‐bound C3b molecules (not depicted here), which are required to strengthen the affinity for C5. Hypothetical model of C3bBb (surface representation, C3b in gray, Bb in orange) bound to substrate C5 (light green, C5d domain in dark green). C3bBb is derived from the dimeric C3bBb‐SCIN complex (PDB 2WIN Rooijakkers et al, 2009), and C5 is modeled based on superposition of the CVF‐C5 complex (PDB 3PVM Laursen et al, 2011) on the C3b molecule from C3bBb. The right panel shows C3bBb bound to C5b (light green, C5d in dark green). The structure of C5b is derived from the structure of the C5b6 complex (PDB 4A5W Hadders et al, 2012) and superimposed on C5 from the model in the left panel.

Superposition of C5b with the C5d domain in the pC5b6 (dark green) and C3b‐like (light blue) orientation. The C3b‐like conformation of C5d was generated based on superposition of the C5d structure (extracted from the pC5b6 structure, PDB 4A5W) on the C3d domain of the second C3b subunit from the dimeric C3bBb‐SCIN structure (PDB 2WIN).

Hypothetical structural models for C5b6 assembly by convertases. (I) Model of pC5b6 bound to C3bBb, as in (A, right). (II) Model of pC5b6, with C5d‐C6 superimposed on C5d in the C3b‐like orientation, as in (B). Note that this orientation allows C6 to extend further toward the membrane relative to the convertase. (III) Model in which C5b6 has dissociated from C3bBb, but adopted the orientation shown in (II). All structural models and superpositions were generated using UCSF Chimera (Pettersen et al, 2004).


Membrane Operations in Molecular Separations

2.12.6 Concluding Remarks

The membrane permeability value can be increased by increasing either the distribution coefficient or the diffusivity for the transported solute. The idea of using thin organic liquid layers as PV membranes seems to be very attractive from this point of view simply because the value of diffusivity in liquids is at least three to four orders of magnitude higher than values in solid polymers and in inorganic membranes. Besides, it is possible to dissolve some selective carriers in the immobilized liquid, so that they will be able to interact with the transported species, increasing the affinity of the solute and, thus, the process selectivity.

There are two primary constraints associated with the use of SLMs. Solvent loss can occur by evaporation, dissolution, or large pressure differences forcing the solvent out of the pore support structure. In addition, carrier loss can occur. This loss can be due to irreversible side reactions or solvent condensation on one side of the membrane. Pressure differences can force the liquid to flow through the pore structure and leach out the carrier. In comparison to solid membranes for PV, LMs are used at moderate temperatures, because the stability problems increase at high temperatures. The stability of LMs can be increased drastically by placing a thin polymer layer on top of the LM however, often it implies a reduction of the permeation flux.

Since ILs do not show measurable vapor pressures, they might overcome stability problems of common SLMs caused by evaporation of the membrane phase. In addition, mechanical stability of the SLM is increased due to the improved wetting properties of ILs.

Main applications reported in literature have been addressed to separate volatile fermentation products and other VOCs from their dilute aqueous solutions. The use of PV coupled to a fermenter acts not only as a means of separation, but also as a production enhancer by reducing product inhibition. Further applications (i.e., organic–organic separations) should be explored taking into account the use of suitable ILs as immobilized liquid in SLMs. ILs can dissolve a very broad spectrum of organic compounds, and their miscibility with these substances can be fine-tuned by changing the nature of the cation and/or anion. However, so far the separations of organic–organic mixtures by using hydrophilic ILs have been limited by interfacial instability phenomena at the feed/membrane interface. Future efforts should be directed toward developing systems with improved stability.


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Beetroot cell membrane permeability experiment - Research Papers - Ptardag1 www.studymode.com/essays/Beetroot-Cell-Membrane-Permeability. 網頁紀錄 - 更多此站結果 Beetroot cell membrane permeability experiment Osmosis, Cell membrane, Semipermeable membrane By Ptardag1 Feb 22, 2003 1408 Words 40481 Views Practical.

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LECTURE PRESENTATIONS For CAMPBELL BIOLOGY, NINTH EDITION Jane B. Reece, Lisa A. Urry, Michael L. Cain, Steven A. Wasserman, Peter V. Minorsky, Robert B. Jackson Chapter 1 Introduction: Themes in the Study of Life Lectures by Erin Barley Kathleen Fitzpatrick © 2011 Pearson Education.


A rapid cell membrane permeability test using flourescent dyes and flow cytometry

A reliable and rapid test to detect cytotoxic chemicals which affect cell membranes is described. Fluorescein diacetate freely penetrates intact cells where it is hydrolyzed to its fluorochrome, fluorescein, which is retained in the cell due to its polarity. On the other hand, ethidium bromide is known to be excluded from the intact cell, staining only nucleic acids of membrane-damaged cells. The combination of both fluorochromes results in counter-staining: intact cells fluoresce green (cytoplasm) and membrane-damaged cells fluoresce red (nucleus and RNA). Rat thymocytes freshly isolated without enzyme treatment were incubated simultaneously with test substance and dye solution fluorescein diacetate and ethidium bromide. A two-parameter analysis was performed on a flow cytometer with an on-line computer. Concentration-dependent effects of various detergents and solvents were quantified by measuring the amount of dye retention, i.e., the decrease or increase in fluorescein—fluorescence (peak shift), and the decrease in dye exclusion (increase in ethidium bromide-staining) relative to the untreated control. The assay can be used for rapid monitoring of chemical insults to cell membranes which precede the decrease of the viability measured by pure dye exclusion techniques.

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ADME-Tox Approaches

5.28.6.2.4 Nonpolar surface area

The Caco-2 cell membrane permeability of three series of peptides and endothelin antagonists could be predicted by a theoretical model that considered both the polar (PSA d) and nonpolar (NPSAd) parts of the dynamic molecular surface area of the molecules. 97,131 The three peptide series were AcHN-X-phenethylamides, AcHN-XD-Phe-NHMe derivatives and d -Phe-oligomers. Experimental log D (octanol/water) values gave a rank order of permeability within each series, but failed across the three series. Possibly, some of the compounds are substrates for one or more transporters present in Caco-2 cells, but this needs further investigation. A strong correlation was found between log D and NPSAd ( r 2 = 0.96 ). A good sigmoidal correlation was obtained when Papp (Caco-2 permeability) was plotted against a linear combination of PSAd and NPSAd. Thus, this model predicts permeability based on a combination of hydrogen-bonding capacity and hydrophobicity. The latter is thought to be related to the transport of a compound from the aqueous environment into the polar head group region of the membrane, while hydrogen bonding is detrimental to transport into the nonpolar interior of the membrane. 131


Getting across the cell membrane: an overview for small molecules, peptides, and proteins

The ability to efficiently access cytosolic proteins is desired in both biological research and medicine. However, targeting intracellular proteins is often challenging, because to reach the cytosol, exogenous molecules must first traverse the cell membrane. This review provides a broad overview of how certain molecules are thought to cross this barrier, and what kinds of approaches are being made to enhance the intracellular delivery of those that are impermeable. We first discuss rules that govern the passive permeability of small molecules across the lipid membrane, and mechanisms of membrane transport that have evolved in nature for certain metabolites, peptides, and proteins. Then, we introduce design strategies that have emerged in the development of small molecules and peptides with improved permeability. Finally, intracellular delivery systems that have been engineered for protein payloads are surveyed. Viewpoints from varying disciplines have been brought together to provide a cohesive overview of how the membrane barrier is being overcome.

Figures

Possible routes of cytosolic entry.…

Possible routes of cytosolic entry. Molecules may passively diffuse across the cell membrane,…

( a ) Cyclosporin A (CsA) in its closed conformation in nonpolar solvent…


Watch the video: PAG The effect of temperature on membrane permeability (December 2022).