What is the thickness of the membrane if only alpha helixes are embedded of a transmembrane protien?

What is the thickness of the membrane if only alpha helixes are embedded of a transmembrane protien?

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Given is the representation of a transmembrane protein. Calculate the thickness of the membrane if only alpha helixes are embedded in it. One turn = 5.4Å

Please read: The reason I didn't submit my attempt is that, I thought because there isn't an exact and precise depiction of the protien molecule and also the question is vague so I hesitated. I counted the number of turns ie 7 but still there was a lingering doubt "what if I am wrong, I would make a fool of myself."

What I know:

  1. Protiens basically bunch of amino acids bound by polypeptide bond.
  2. Primary structure is the sequence of the amino acids.
  3. The secondary structure us due to the Hydrogen Bonding between the Oxygen atoms having partial -ve charge and Hydrogen atom attached to the Nitrogen, within the backbone only. eg: alpha-helixes, beta-sheets, Turns, Loops
  4. Tertiary Structure is due to the interaction between the different R groups in the amino acids. Like the disulphide bonds, hydrophobic interactions, H-bonds, Vanderwaal forces.
  5. Quaternary Structure is due to the overall 3D structure which is result of the aggregation of the polypeptide subunits.

Please go easy on me, its my first time posting here and I am still a 1st year in college and 2nd Sem is gonna start next week.

The question begins by telling you that only the alpha helices are embedded in the membrane. This means that if you can calculate how long the alpha helices are, you can work out the thickness of the membrane it is embedded in.

Alpha helices look like spirals (similar to fusilli pasta shapes). These are highlighted in figure A) below.

The question also tells you that each turn of the alpha helix is 5.4 angstroms. One turn if shown in figure B) below.

Looking at the picture, it seems that each helix is about 7 turns long. As each turn is 5.4 A, the length of the helices must be approximately 7 * 5.4 A which is 37.8 A.

Therefore, using the information that the membrane must be about as thick as the helices in the transmembrane proteins are long, you can deduce that the membrane must be up to 37.8 A thick, however the membrane may not actually be this thick as the question from the paper doesn't specify how much of the alpha helices are embedded in the membrane (60%, 80%, 100%?). I guess the assumption is that the entire alpha helix section of the transmembrane protein is fully embedded within the membrane.

Lipid-protein interactions in biological membranes: a structural perspective

Lipid molecules bound to membrane proteins are resolved in some high-resolution structures of membrane proteins. An analysis of these structures provides a framework within which to analyse the nature of lipid-protein interactions within membranes. Membrane proteins are surrounded by a shell or annulus of lipid molecules, equivalent to the solvent layer surrounding a water-soluble protein. The lipid bilayer extends right up to the membrane protein, with a uniform thickness around the protein. The surface of a membrane protein contains many shallow grooves and protrusions to which the fatty acyl chains of the surrounding lipids conform to provide tight packing into the membrane. An individual lipid molecule will remain in the annular shell around a protein for only a short period of time. Binding to the annular shell shows relatively little structural specificity. As well as the annular lipid, there is evidence for other lipid molecules bound between the transmembrane alpha-helices of the protein these lipids are referred to as non-annular lipids. The average thickness of the hydrophobic domain of a membrane protein is about 29 A, with a few proteins having significantly smaller or greater thicknesses than the average. Hydrophobic mismatch between a membrane protein and the surrounding lipid bilayer generally leads to only small changes in membrane thickness. Possible adaptations in the protein to minimise mismatch include tilting of the helices and rotation of side chains at the ends of the helices. Packing of transmembrane alpha-helices is dependent on the chain length of the surrounding phospholipids. The function of membrane proteins is dependent on the thickness of the surrounding lipid bilayer, sometimes on the presence of specific, usually anionic, phospholipids, and sometimes on the phase of the phospholipid.

Types of Membrane Protein

Based on location and structure, the membrane proteins are typically classified into the following groups:

Integral Proteins

They refer to the intrinsic proteins, which completely spans the phospholipid bilayer membrane. Transmembrane or integral proteins have domains within the cytoplasm and towards the extracellular ends of the lipid bilayer.

They either have multiple spanning segments and contribute about 25-30% of all the encoded proteins. Integral proteins are amphipathic in nature, comprising both hydrophilic and hydrophobic regions.

The intermediate portion of the phospholipid bilayer shows a hydrophobic character, while the rest of the part (protruded outwards) shows hydrophilic behaviour. Due to membrane fluidity, they move laterally within the biological cell membrane.

They are firmly attached to the cell membrane. Thus, the integral proteins are not easily separable and require ionic and non-ionic detergent treatment (like SDS and Triton X-100). SDS denatures the structure of proteins, while Triton X-100 does not alter the protein conformation.

Therefore, the solubilisation of integral proteins by the detergents helps us to study the composition of amino acids, molecular weight and other physicochemical properties. The integral proteins can be monotopic and polytopic, depending upon the number of alpha-helices.

  • Monotopic integral proteins possess a single coiled alpha helix.
  • Polytopic integral proteins possess combinations of coiled alpha-helices.

Not all the transmembrane proteins have alpha-helices only a few have beta-barrel sheets. Glycophorin-A is the best example of an integral protein found in erythrocytes comprising 131 amino acid residues and primarily glycoproteins.

The transmembrane proteins have alpha-helices, which generally contain 21-26 hydrophobic amino acid residues. They undergo coiling to form alpha-helix and facilitate the spanning of the bilayer membrane.

Few transmembrane proteins consist of antiparallel arranged beta-strands, known as beta-barrels. Porin beta-barrels contain 16 stranded antiparallel beta-sheets with a highly hydrophilic interior and hydrophobic exterior.

Peripheral Proteins

They refer to the extrinsic proteins, which associate with the lipid bilayer through weak electrostatic and hydrogen bond interactions. Peripheral proteins are easily separable under the exposure of high pH and treatment with high salt concentration.

They are located directly on the polar heads of the phospholipid bilayer membrane or indirectly on the transmembrane channels. The peripheral proteins help in anchoring, cell support, and transmission of transmembrane signals.

It does not interface with the cell membrane’s hydrophobic core. Peripheral proteins are water-soluble and present higher in number than integral proteins. Extrinsic proteins are less mobile.

Lipid Anchored Proteins

They refer to the lipid-linked proteins, which binds covalently to the lipid membrane either through a fatty acid chain, prenyl group or often via an oligosaccharide complex. The peripheral proteins attach with the lipid membrane through glycosylphosphatidylinositol linkage to constitute “GPI- anchored proteins”.

Lipid-linked proteins have the characteristic property that they are located on either side of the biological cell membrane. They belong to the class of “Proteolipids”.

Prenylated, fatty acylated, and glycosylphosphatidylinositol linked proteins are the three distinct kinds of lipid anchored proteins. Lipid-anchored proteins possess multiple lipid groups and serve as the hydrolytic enzymes, adhesion molecules, receptor proteins, etc.

Functions of Membrane Proteins

The lipid content of the cell membrane helps in forming a semipermeable barrier between the surrounding and protoplasm. Thus, transport proteins allow the influx and efflux of particular solutes.

Some transport proteins actively shuttle the various ions and molecules across the selective barrier by changing their conformation and hydrolyzing a high energy molecule ATP.

Membrane proteins also possess enzyme complexes, which carry out the sequential steps during different metabolic pathways. They also mediate signal transduction or signal-relaying by the specific binding between the chemical messengers with the binding site of the membrane proteins.

Glycoprotein-channels facilitate cell to cell recognition by specific binding between the receptors and ligands of the adjacent cells. The membrane proteins join at different gap junctions or show inter-cellular joining, which aids in cell-cell communication.

The cytoskeleton forms the interlinking protein filaments or microfilaments associated with the transport proteins, which promote cell shape and coordination between the cells doing extracellular and intracellular activities.

What is the thickness of the membrane if only alpha helixes are embedded of a transmembrane protien? - Biology

Figure 1: An electron micrograph of an E. coli cell highlighting the width of the cell inner and outer membranes and the cell wall. Zoom in: a schematic of the lipid bilayer. The red circle denotes the hydrophilic head consisting of a polar phosphoglycerol group and the pink lines represent the hydrocarbon chains forming a tight hydrophobic barrier excluding water as well as polar or charged compounds. Two tails are drawn per head but there could also be three or four. (Electron microscopy image adapted from A. Briegel et al. Proc. Nat. Acad. Sci., 106:17181, 2009.)

One of the key defining characteristics of living organisms is that cells are separated from their external environment by a thin, but highly complex and heterogeneous cell membrane. These membranes can come in all sorts of shapes and molecular compositions, though generally they share the property of being made up of a host of different lipid molecules and that they are riddled with membrane proteins. Indeed, if we take the mass of all the proteins that are present in such a membrane and compare it to the mass of all of the lipids in the same membrane, this so-called protein-to-lipid mass ratio is often greater than one (BNID 105818). This assertion applies not only to the plasma membranes that separate the cellular contents from the external world, but also to the many organellar membranes that are one of the defining characteristics of eukaryotic cells.

The thickness of this crucial but very thin layer in comparison to the diameter of the cell, is similar to the thickness of an airplane fuselage in comparison with the plane’s body diameter. The key point of this analogy is simply to convey a geometric impression of the thickness of the membrane relative to the dimensions of the cell using familiar everyday objects. In the case of an airplane, the thickness of the exterior shell is roughly 1 cm in comparison with the overall diameter of roughly 5 m, resulting in an aspect ratio of 1:500. How can we estimate the aspect ratio for the biological case? With a few exceptions, such as in Archaea, the lipid part of the cell membrane is a bilayer of lipids with the tails on opposite leaflets facing each other (see Figure 1). These membranes spontaneously form as a relatively impermeable and self-mending barrier at the cell’s (or organelle’s) periphery as discussed in the section on the cell’s membrane permeability. The length scale of such structures is given by the lipid molecules themselves as shown in Figure 2. For example the prototypical phospholipid dipalmitoyl- phosphatidylcholine, has a head to tail length of 2 nm (BNID 107241, 107242). This implies an overall bilayer membrane thickness of 4 nm (3 nm of which are strongly hydrophobic and the rest being composed of the polar heads, (BNID 107247)). For a 2 micron cell diameter (a relatively large bacterium or a very small eukaryotic cell), the 4 nm thickness implies an aspect ratio of 1:500, similar to the case of an airplane. Larger numbers are sometimes quoted probably resulting from the effective increase due to proteins and lipopolysaccharides sticking out of the membrane. For example, the lipopolysaccharide incorporated in the Gram-negative bacterial outer membrane nearly doubles the diameter of the cell.

Figure 2: Characteristic relative sizes and shapes of the lipid molecules making up biological membranes.

The story of how lipid size was initially estimated has a long and interesting history as vividly described in Charles Tanford’s little book “Ben Franklin Stilled the Waves”. Specifically, the story begins with experiments of Benjamin Franklin who explored the capacity of oils to still the waves. Franklin performed his experiments in a pond near London and said of them, “the oil, though not more than a teaspoonful, produced an instant calm over a space several yards square, which spread amazingly and extended itself gradually until it reached the leeside, making all that quarter of the pond, perhaps half an acre, as smooth as a looking glass.” The calming of the waves is attributed to a monolayer of oil forming on the surface of the water and causing damping through energy dissipation. A similar approach to calming waves was taken by sailors at the time of the Romans by dumping oil (such as whalers using blubber) in rough seas. Energy is dissipated as the oil film flows and gets compressed and dilated during the movement of the waves. Using Franklin’s own dimensions for the size of his oil slick (i.e. ½ acre ≈ 2000 m 2 ) and the knowledge of the initial teaspoon volume (i.e. 1 teaspoon ≈ 5 cm 3 ), we see that his oil formed a single layer with a thickness of several nanometers. To be precise, using the numbers above one finds a thickness of roughly 2.5 nm. More precise measurements were undertaken by Agnes Pockels, who invented an experimental technique used to construct lipid monolayers that made it possible to settle the question of molecular dimensions precisely. Lord Rayleigh performed small-scale versions of the Franklin experiment in an apparatus similar to what is now known as the “Langmuir trough” and permits spreading of a monolayer of molecules on a liquid surface and detecting their presence with a small wire that squeezes this monolayer.

Each layer of the cell membrane is made up of molecules similar in character to those investigated by Franklin, Rayleigh and others. In particular, the cell membrane is composed of phospholipids which contain a head group and a fatty acid tail which is roughly 10-20 carbons long. An average carbon-carbon bond length projected on the chain and thus accounting for the tail’s zigzag shape arising from carbon’s tetrahedral orbital shape is lcc=0.126 nm (BNID 109594). The overall tail length is nc x lcc where nc is the number of carbon atoms along the chain length. Overall the two tails end-to-end plus the phosphoglycerol head groups have a length of ≈4nm (BNID 105821, 100015, 105297 and 105298).

Figure 3: The membrane with some notable constituents. The extent of protrusion of proteins from the cell membrane is evident. The fraction of membrane surface occupied by proteins in this cross section depiction is similar to that actually found in cells. (Courtesy of David Goodsell)

Unsurprisingly, membrane proteins are roughly as thick as the membranes they occupy. Many membrane proteins like ion channels and pumps are characterized by transmembrane helices that are ≈4 nm long, and have physicochemical properties like that of the lipids they are embedded in. Often these proteins also have regions which extend into the space on either side of the membrane. This added layer of protein and carbohydrate fuzz adds to the “thickness” of the membrane. This is evident in Figure 3 where some of the membrane associated proteins are shown to scale in cross section. Due to these extra constituents that also include lipopolysaccharides, the overall membrane width is variably reported to be anywhere between 4 and 10 nm. The value of 4 nm is most representative of the membrane shaved off from its outer and inner protrusions. This value is quite constant across different organellar membranes as shown recently for rat hepatocyte via x-ray scattering where the ER, Golgi, basolateral and apical plasma membranes, were 3.75±0.04 nm, 3.95±0.04 nm, 3.56±0.06 nm, and 4.25±0.03 nm, respectively (BNID 105819, 105820, 105822, 105821). We conclude by noting that the cell membrane area is about half protein (BNID 106255) and the biology and physics of the dynamics taking place there is still intensively studied and possibly holds the key to the action of many future drugs.

Structure-based statistical analysis of transmembrane helices

Recent advances in determination of the high-resolution structure of membrane proteins now enable analysis of the main features of amino acids in transmembrane (TM) segments in comparison with amino acids in water-soluble helices. In this work, we conducted a large-scale analysis of the prevalent locations of amino acids by using a data set of 170 structures of integral membrane proteins obtained from the MPtopo database and 930 structures of water-soluble helical proteins obtained from the protein data bank. Large hydrophobic amino acids (Leu, Val, Ile, and Phe) plus Gly were clearly prevalent in TM helices whereas polar amino acids (Glu, Lys, Asp, Arg, and Gln) were less frequent in this type of helix. The distribution of amino acids along TM helices was also examined. As expected, hydrophobic and slightly polar amino acids are commonly found in the hydrophobic core of the membrane whereas aromatic (Trp and Tyr), Pro, and the hydrophilic amino acids (Asn, His, and Gln) occur more frequently in the interface regions. Charged amino acids are also statistically prevalent outside the hydrophobic core of the membrane, and whereas acidic amino acids are frequently found at both cytoplasmic and extra-cytoplasmic interfaces, basic amino acids cluster at the cytoplasmic interface. These results strongly support the experimentally demonstrated biased distribution of positively charged amino acids (that is, the so-called the positive-inside rule) with structural data.

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How proteins become embedded in a cell membrane

The experiment: a tiny cantilever measuring just a few nanometers in thickness is used to isolate a membrane protein (red) and transport it to a helper protein (purple) in a different location. Credit: Visualisations: Serdiuk T et al., Science Advances 2019, edited

Many proteins with important biological functions are embedded in a biomembrane in the cells of humans and other living organisms. But how do they get in there in the first place? Researchers in the Department of Biosystems Science and Engineering at ETH Zurich have investigated the matter.

Nearly one-third of all proteins in living beings are firmly embedded in a biomembrane—either in a cell's outer membrane or in the boundaries of internal cellular compartments. There, these membrane proteins perform important tasks, serving, for instance, as molecular channels for transporting metabolites and nutrients through the membrane or as sensor proteins for sensing the cellular environment.

A team of researchers led by Daniel J. Müller, a professor in the Department of Biosystems Science and Engineering at ETH Zurich in Basel, has now investigated how membrane proteins manage to get into the membranes. To do this, they used a highly precise method that enables them to extract individual proteins from, or deposit them on, membranes. This method, known as single-molecule force spectroscopy, lets scientists guide a computer-controlled cantilever measuring just a few nanometers in thickness to a specific location on a membrane's surface with utmost precision. Molecular adhesive forces cause a protein located there to adhere to the cantilever.

Role of two helper proteins

In experiments with bacterial proteins, the researchers were able to clarify the role of two helper proteins—an insertase and a translocase—that enable the membrane proteins to embed themselves in the membrane. Insertase is a single protein, while translocase is a complex composed of multiple proteins. Both of them ensure that a pore opens up in the membrane. "In the case of insertase, we can think of this pore as a slide. The membrane protein is initially present as an unstructured peptide strand that slips down this slide into the membrane. In the membrane, this peptide strand then takes on its functional three-dimensional shape," explains ETH Professor Müller. "Once the membrane protein has successfully become three-dimensional and embedded itself in the membrane, the helper protein detaches and forms a slide at a different location in the membrane for the next protein," he continues.

The helper protein (here a translocase, purple) is already located in the biomembrane (grey) and helps a membrane protein in the form a peptide strand (left, red) embed itself in the membrane (right). Credit: Visualisations: Serdiuk T et al., Science Advances 2019

Up to now, research into how these helper proteins function was imprecise and used only short peptides or was conducted only outside of biomembranes. "We have now observed and described for the first time, step by step, how an entire protein embeds itself in a membrane and takes on a three-dimensional form," says Tetiana Serdiuk, a postdoc in ETH Professor Müller's group and the study's first author.

The ETH researchers were also able to show the differences in how insertases and translocases work: insertases insert peptide strands into the membrane relatively quickly but clumsily. "This means they work well, particularly with small proteins," says Müller. Translocases, on the other hand, insert peptide strands into the membrane section by section, making them better suited for more complex proteins.

Important for medicine

This study is a case of classic basic research, which is particularly significant in view of the importance of membrane proteins to medicine, as Müller emphasises: "Around half of all drugs act on membrane proteins, and we need to understand how these membrane proteins form and how they work."

In addition, single-molecule force spectroscopy, which the ETH scientists further refined for this study, could be used in other applications: in connection with the National Center of Competence in Research (NCCR) for Molecular Systems Engineering, Müller and other scientists are working to develop artificial biological cells. "This method could be used to custom-fit biomembranes with proteins, essentially programming them," says the ETH professor. "Artificial cells of this kind could one day be used as molecular factories to produce pharmaceuticals on an industrial scale."

G protein-coupled receptor

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G protein-coupled receptor (GPCR), also called seven-transmembrane receptor or heptahelical receptor, protein located in the cell membrane that binds extracellular substances and transmits signals from these substances to an intracellular molecule called a G protein (guanine nucleotide-binding protein). GPCRs are found in the cell membranes of a wide range of organisms, including mammals, plants, microorganisms, and invertebrates. There are numerous different types of GPCRs—some 1,000 types are encoded by the human genome alone—and as a group they respond to a diverse range of substances, including light, hormones, amines, neurotransmitters, and lipids. Some examples of GPCRs include beta-adrenergic receptors, which bind epinephrine prostaglandin E2 receptors, which bind inflammatory substances called prostaglandins and rhodopsin, which contains a photoreactive chemical called retinal that responds to light signals received by rod cells in the eye. The existence of GPCRs was demonstrated in the 1970s by American physician and molecular biologist Robert J. Lefkowitz. Lefkowitz shared the 2012 Nobel Prize for Chemistry with his colleague Brian K. Kobilka, who helped to elucidate GPCR structure and function.

A GPCR is made up of a long protein that has three basic regions: an extracellular portion (the N-terminus), an intracellular portion (the C-terminus), and a middle segment containing seven transmembrane domains. Beginning at the N-terminus, this long protein winds up and down through the cell membrane, with the long middle segment traversing the membrane seven times in a serpentine pattern. The last of the seven domains is connected to the C-terminus. When a GPCR binds a ligand (a molecule that possesses an affinity for the receptor), the ligand triggers a conformational change in the seven-transmembrane region of the receptor. This activates the C-terminus, which then recruits a substance that in turn activates the G protein associated with the GPCR. Activation of the G protein initiates a series of intracellular reactions that end ultimately in the generation of some effect, such as increased heart rate in response to epinephrine or changes in vision in response to dim light (see second messenger).

Both inborn and acquired mutations in genes encoding GPCRs can give rise to disease in humans. For example, an inborn mutation of rhodopsin results in continuous activation of intracellular signaling molecules, which causes congenital night blindness. In addition, acquired mutations in certain GPCRs cause abnormal increases in receptor activity and expression in cell membranes, which can give rise to cancer. Because GPCRs play specific roles in human disease, they have provided useful targets for drug development. The antipsychotic agents clozapine and olanzapine block specific GPCRs that normally bind dopamine or serotonin. By blocking the receptors, these drugs disrupt the neural pathways that give rise to symptoms of schizophrenia. There also exist a variety of agents that stimulate GPCR activity. The drugs salmeterol and albuterol, which bind to and activate beta-adrenergic GPCRs, stimulate airway opening in the lungs and thus are used in the treatment of some respiratory conditions, including chronic obstructive pulmonary disease and asthma.

Plasma membranes enclose and define the borders between the inside and the outside of cells. They are typically composed of dynamic bilayers of phospholipids into which various other lipid-soluble molecules and proteins have also been embedded. These bilayers are asymmetric&mdashthe outer leaf differing from the inner leaf in lipid composition and in the proteins and carbohydrates that are displayed to either the inside or outside of the cell. One major function of the outer cell membrane is to communicate the cell&rsquos unique identity to other cells. The proteins, lipids, and sugars displayed on the cell membrane allow for cells to be detected by and to interact with specific partners.

Various factors influence fluidity, permeability, and various other physical properties of the membrane. These include temperature, the configuration of the fatty acid tails (some

by double bonds), sterols (i.e., cholesterol) embedded in the membrane, and the mosaic nature of the many proteins embedded within it. The plasma membrane is &ldquoselectively permeable&rdquo. This means it allows only some substances through while excluding others. In addition, the plasma membrane must, sometimes, be flexible enough to allow certain cells, such as amoebae, to change shape and direction as they move through the environment, hunting smaller, single-celled organisms.

Cellular membranes

A subgoal in our "build-a-cell" design challenge is to create a boundary that separates the "inside" of the cell from the environment "outside". This boundary needs to serve multiple functions that include:

  1. Act as a barrier by blocking some compounds from moving in and out of the cell.
  2. Be selectively permeable

Figure 1. The diameter of a typical balloon is 25cm and the thickness of the plastic of the balloon of around 0.25mm. This is a 1000X difference. A typical eukaryotic cell will have a cell diameter of about

and a cell membrane thickness of 5nm. This is a 10,000X difference.

Fluid mosaic model

The fluid mosaic model describes the dynamic movement of the

proteins, sugars, and lipids embedded in the cell&rsquos plasma membrane.

For some insight into the history of our understanding of the plasma membrane structure, click here.

It is sometimes useful to start our discussion by recalling the size of the cell membrane relative to the size of the entire cell

. Plasma membranes range from 5 to 10

in thickness. For comparison, human red blood cells, visible via light microscopy, are approximately 8

wide, or approximately 1,000 times wider than a plasma membrane is thick. This means that the cellular barrier is very thin compared to the size of the volume it encloses. Despite this dramatic size differential, the cellular membrane must

still carry out its key barrier, transport and cellular recognition capacities and so must be a relatively &ldquosophisticated&rdquo and dynamic structure.

Figure 2. The fluid mosaic model of the plasma membrane describes the plasma membrane as a fluid combination of phospholipids, cholesterol, and proteins. Carbohydrates attached to lipids (glycolipids) and

proteins (glycoproteins) extend from the outward-facing surface of the membrane.

The principal components of a plasma membrane are lipids (phospholipids and cholesterol), proteins, and carbohydrates. Carbohydrates are present only on the exterior surface of the plasma membrane and

to proteins, forming glycoproteins, or

lipids, forming glycolipids. The proportions of proteins, lipids, and carbohydrates in the plasma membrane may vary with organism and cell type. In a typical human cell, proteins account for a massive 50 percent of the composition by mass, lipids (of all types) account for about 40 percent of the composition by mass, and carbohydrates account for the remaining 10 percent of the composition by mass. However, cellular functional specialization may cause these ratios of components to vary dramatically. For example, myelin, an outgrowth of the membrane of specialized cells, insulates the axons of the peripheral nerves, contains only 18 percent protein and 76 percent lipid.

mitochondrial inner membrane contains 76 percent protein and only 24 percent lipid and the plasma membrane of human red blood cells is 30 percent lipid.


Phospholipids are major constituents of the cell membrane.

of a glycerol backbone to which

fatty acid tails and a phosphate group

- one to each of each of the glycerol carbons atoms. The phospholipid is therefore an amphipathic molecule, meaning it has a hydrophobic part (fatty acid tails) and a hydrophilic part (phosphate head group).

Make sure to note in Figure 3 that the phosphate group has an R group linked to one of the oxygen atoms. R is a variable commonly used in these types of diagrams to indicate that some other atom or molecule is bound at that position. That part of the molecule can be different in different phospholipids&mdashand will impart some different chemistry to the whole molecule. At the moment, however, you are responsible for being able to recognize this type of molecule (no matter what the R group is) because of the common core elements&mdashthe glycerol backbone, the phosphate group, and the two hydrocarbon tails.

Figure 3. A phospholipid is a molecule with two fatty acids and a

phosphate group attached to a glycerol backbone.

The phosphate may be modified by the addition of charged or polar chemical groups

. Several chemical R groups may

the phosphate. Choline, serine, and ethanolamine

here. These attach to the phosphate group at the position labeled R via their hydroxyl groups.

to an aqueous environment, they can spontaneously arrange themselves into various structures including micelles and phospholipid bilayers. The latter is the basic structure of the cell membrane. In a phospholipid bilayer, the phospholipids associate with one another into two

sheets. In each sheet nonpolar parts of the phospholipids face inward towards one another, composing the internal part of the membrane, and polar head groups facing oppositely to both the aqueous extracellular and intracellular environments.

Possible NB Discussion Point

Earlier in the course, we discussed the Second Law of Thermodynamics, which states that the overall entropy of the universe is always increasing. Apply this law in the context of the formation of the lipid bilayer membrane. How is it possible that the lipids are able to spontaneously arrange themselves into such an organized structure instead of scatter into a more disordered state? Or in other words -- if the second law holds true, then how exactly does the spontaneous lipid organization lead to increased entropy?

Figure 4. In the presence of water, some phospholipids will spontaneously arrange themselves into a micelle.

The lipids will be arranged

such that their polar groups will be on the outside of the micelle, and the nonpolar tails will be on the inside. A lipid bilayer can also form, a two layered sheet only a few nanometers thick. The lipid bilayer

two layers of phospholipids organized in a way that all the hydrophobic tails align side by side in the center of the bilayer and

by the hydrophilic head groups.
Source: Created by

Membrane proteins

Proteins make up the second major component of plasma membranes. Integral membrane proteins, as their name suggests, integrate completely into the membrane structure, and their hydrophobic membrane-spanning regions interact with the hydrophobic region of the phospholipid bilayer.

Some membrane proteins associate with only one half of the bilayer, while others stretch from one side of the membrane to the other, and

to the environment on either side. Integral membrane proteins may have one or more transmembrane segments typically

20&ndash25 amino acids. Within the transmembrane segments, hydrophobic amino acid variable groups arrange themselves to form a chemically complementary surface to the hydrophobic tails of the membrane lipids.

Peripheral proteins are found on only one side of the membrane, but never embed into the membrane. They can be on the intracellular side or the extracellular side, and weakly or temporarily associated with the membranes.

Figure 5. Integral membranes proteins may have one or more &alpha -helices (pink cylinders) that span the membrane (examples 1 and 2), or they may have &beta-sheets (blue rectangles) that span the membrane (example 3). (credit: &ldquoFoobar&rdquo/Wikimedia Commons)


Carbohydrates are a third major component of plasma membranes. They are always found on the exterior surface of cells and are bound either to proteins (forming glycoproteins) or to lipids (forming glycolipids). These carbohydrate chains may consist of 2&ndash60 monosaccharide units and can be either straight or branched. Along with peripheral proteins, carbohydrates form specialized sites on the cell surface that allow cells to recognize each other (one of the core functional requirements noted above.

Membrane fluidity

The integral proteins and lipids exist in the membrane as separate molecules and they "float" in the membrane, moving with respect to one another. The membrane is not like a balloon, however because of the elastic properties of its plastic a balloon can easily grow and shrink its surface area without popping and while also maintaining the same rough circular shape. By contrast the plasma membrane is not able to withstand isotropic stretching or compression and can be easily popped when an imbalance of solute between inside and out causes water to rush in suddenly. A sudden loss of water will cause it to shrivel and wrinkle, dramatically changing the shape of the cell. it is fairly rigid and can burst if penetrated or if a cell takes in too much water and the membrane is stretched too far. However, because of its mosaic nature, a very fine needle can easily penetrate a plasma membrane without causing it to burst (the lipids flow around the needle point), and the membrane will self-seal when the needle is extracted.

Different organisms and cell types in multicellular organisms can tune fluidity of their membrane to be more compatible with specialized functions and/or in response to environmental factors. This tuning can be accomplished by adjusting the type and concentration of various components of the membrane, including the lipids, their degree of saturation, the lipids, their degree of saturation, the proteins, and other molecules like cholesterol. There are two other factors that help maintain this fluid characteristic. One factor is the nature of the phospholipids themselves. In their saturated form, the fatty acids in phospholipid tails are saturated with hydrogen atoms. There are no double bonds between adjacent carbon atoms, resulting in tails that are relatively straight. By contrast, unsaturated fatty acids do not have a full complement of hydrogen atoms on their fatty acid tails and therefore contain some double bonds between adjacent carbon atoms a double bond results in a bend in the string of carbons of approximately 30 degrees.

Figure 6. Any given cell membrane will be composed of a combination of saturated and unsaturated phospholipids. The ratio of the two will influence the permeability and fluidity of the membrane. A membrane composed of completely saturated lipids will be dense and less fluid, and a membrane composed of completely unsaturated lipids will be very loose and very fluid.

Saturated fatty acids, with straight tails, are compressed by decreasing temperatures, and they will press in on each other, making a dense and fairly rigid membrane. Conversely, when unsaturated fatty acids are compressed, the &ldquokinked&rdquo tails elbow adjacent phospholipid molecules away, maintaining some space between the phospholipid molecules. This &ldquoelbow room&rdquo helps to maintain fluidity in the membrane at temperatures at which membranes with high concentrations of saturated fatty acid tails would &ldquofreeze&rdquo or solidify. The relative fluidity of the membrane is particularly important in a cold environment. Many organisms (fish are one example) are capable of adapting to cold environments by changing the proportion of unsaturated fatty acids in their membranes in response to the lowering of the temperature.


Animal cells have cholesterol, an additional membrane constituent that assists in maintaining fluidity. Cholesterol, which lies right in between the phospholipids in the membrane, tends to dampen the effects of temperature on the membrane.Cholesterol both stiffens and increases membrane fluidity, depending on the temperature. Low temperatures cause phospholipids to pack together more tightly, creating a stiffer membrane. In this case, the cholesterol molecules serve to space the phospholipids apart and prevent the membrane from becoming totally rigid. Conversely, higher temperatures contribute to phospholipids moving farther apart from each other and therefore a more fluid membrane, but cholesterol molecules in the membrane take up space and prevent the complete dissociation of phospholipids.

Thus, cholesterol extends, in both directions, the range of temperature in which the membrane is appropriately fluid and consequently functional. Cholesterol also serves other functions, such as organizing clusters of transmembrane proteins into lipid rafts.

Figure 7. Cholesterol fits between the phospholipid groups within the membrane.

Review of the components of the membrane

The components and functions of the plasma membrane
Component Location
Phospholipid Main fabric of the membrane
Cholesterol Between phospholipids and between the two phospholipid layers of animal cells
Integral proteins (e.g., integrins) Embedded within the phospholipid layer(s) may or may not penetrate through both layers
Peripheral proteins On the inner or outer surface of the phospholipid bilayer not embedded within the phospholipids
Carbohydrates (components of glycoproteins and glycolipids) Generally attached to proteins on the outside membrane layer

One major difference setting archaea apart from eukaryotes and bacteria is their membrane lipid composition. Though eukaryotes, bacteria, and archaea all use glycerol backbones in their membrane lipids, Archaea use long isoprenoid chains (20-40 carbons in length, derived from the five-carbon lipid isoprene) that are attached via ether linkages to glycerol, whereas eukaryotes and bacteria have fatty acids bonded to glycerol via ester linkages.

The polar head groups differ based on the genus or species of the Archaea and consist of mixtures of glyco groups (mainly disaccharides), and/or phospho groups primarily of phosphoglycerol, phosphoserine, phosphoethanolamine or phosphoinositol. The inherent stability and unique features of archaeal lipids have made them a useful biomarker for archaea within environmental samples, though approaches based on genetic markers are now more commonly used.

A second difference between bacterial and archaeal membranes that is associated with some archaea is the presence of monolayer membranes, as depicted below. Notice that the isoprenoid chain is attached to the glycerol backbones at both ends, forming a single molecule consisting of two polar head groups attached via two isoprenoid chains.

Figure 8. The exterior surface of the archaeal plasma membrane is not identical to the interior surface of the same membrane.

Figure 9. Comparisons of different types of archaeal lipids and bacterial/eukaryotic lipids

Bulk Lipid-Protein Interactions

Effect of Viscosity of the Membrane

The movement of any molecule or part of a molecule inside the membrane is influenced by the fluidity of this environment [9]. Every molecule experiences a frictional drag force during their movement inside the lipid bilayer structure of a biological membrane, and resistance of the fluid to this motion can be expressed in terms of the viscosity. By Stokes&rsquo Law (Equation ef), the rate of particle motion can be expressed as a function of the drag force, membrane viscosity, and particle size [3]. This equation assumes the absence of turbulent flow (a good assumption for the comparatively calm cell membrane).

  • (F): drag force (e.g. Newtons)
  • (mu): membrane viscosity (e.g. centipoise, Pascal*seconds)
  • (R): particle size (e.g. meters, nanometers)
  • (v): speed of particle motion (e.g. meters per second)

Larger particles, more viscous membranes, and more rapid motion thus all result in increased drag force. More viscous membranes can arise from the phase (e.g. liquid disordered, liquid ordered, or gel) of the lipids in the membrane the phase is determined by the number of double bonds in the fatty acid tails of the lipids and the environmental temperature, with fewer double bonds or a temperature below the critical temperature for the lipids resulting in a more viscous membrane.

Movement of a protein in the membrane is dictated by the frictional resistance and the molecular restoring forces acting on it. A fluorescence polarization anisotropy study on the rotation of tryptophan (Trp) residues showed that the motion of Trp was affected by the viscosity of the lipid environment for small scale motion, and the amplitude of this motion increased with the temperature as the viscosity decreased. For example, at a viscosity of 1 centipoise (approximately the viscosity of water at room temperature and on the order of magnitude of the viscosity of the membrane) the authors expect local rotational motion of a given peptide within the protein to be largely determined by the surrounding peptide residues [17].

While the viscosity of the membrane can affect the rate of motion of the protein, the most thermodynamically favorable conformation of the protein should not depend on membrane viscosity [9]. This is because the thermodynamic properties of a system are inherently time-independent, but viscosity is a time-related parameter (since viscosity has units of pressure*time). This means a change in viscosity cannot directly increase or decrease the activation energy required to induce a conformational change--the most stable conformation of a protein is independent of the membrane viscosity. Changes in membrane viscosity can, however, change the amount of time required for the protein to assume this conformation. Additionally, if a change in membrane viscosity is the result of a change in temperature, the rate of protein function will be affected because rate constants are temperature-dependent [9]. This indirect effect can be worthwhile to consider in the study of isolated membrane proteins, which may take place at room temperature rather than at the temperature of the protein's natural environment.

Another important effect of membrane viscosity has also been revealed by molecular dynamics simulations. It has been illustrated that the effect of solvent viscosity on protein motion is important if the protein both directly contacts the solvent and has a rate of motion comparable to the environment. In other words, high frequency motion of a protein that does not overlap the solvent motion does not depend on the solvent&rsquos viscosity [2]. Lipids are important in this context because viscosity of a biological membrane is mainly influenced by the types of lipids present, as the lipid composition is one of the main determinants of the fluidity.

Highly viscous membranes are indicative of increased ordering of the lipid fatty acid tails, often facilitated by interactions with smaller molecules such as sphingomyelin or cholesterol. This can result in increased membrane durability and impermeability. Additionally, the fluidity of the membrane environment surrounding a membrane protein can impact its function. For example, the human epidermal growth factor receptor (EGFR) was found to be non-functional in an environment in which both highly fluid liquid disordered and more viscous liquid ordered domains could exist. However, in a purely liquid disordered environment, EGFR retained its function, revealing a functional dependence on the fluidity of the membrane [3].

Figure (PageIndex<3>): The elasticity of the bilayer is often facilitated by small molecules such as sphingomyelin and cholesterol, and can impact the behavior of membrane proteins [15].

(See the main phase transitions page for more information on membrane fluidity.)

Localized areas of high membrane viscosity can also be indicative of the existence of lipid rafts, which are believed to play a role in protein isolation and function. Lipid rafts are transient sphingomyelin- and cholesterol-rich : (10-200 nm in diameter) within the cell membrane. The local abundance of sphingomyelin and/or cholesterol results in liquid-liquid phase separation, meaning lipid rafts are typically composed of the liquid ordered (Lo) phase. This means the lipids within rafts are more tightly packed and highly ordered than the bulk membrane, leading to increased local rigidity and decreased fluidity [18]. Some proteins can bind glycosphingolipids or sphingomyelin, which are involved in the recruitment of proteins to lipid rafts.

Effect of Membrane Curvature

Dynamic curvature of plasma or intercellular membrane can be dictated by the interactions between proteins and lipids. In fact, cells can employ various mechanisms to sense curvature as a way to create regions of active membrane trafficking. Lipid composition is a major influence on membrane curvature based on their chemical properties and/or the size of their headgroup. Presence of certain lipids are key to interact with certain peripheral membrane proteins in order to induce a necessary curvature. For example, phosphoinositides are required for budding of clathrin-coated vesicles, as the required machinery (e.g. coat proteins) can specifically bind to these lipids. The reason for this is that the headgroups of phosphoinositides are negatively charged, resulting in electrostatic repulsion that can contribute to membrane curvature [14].

Another study illustrated that NSA4(1-48), a key transmembrane protein that mediates replication of Dengue virus (a mosquito born single positive-stranded RNA virus), has an affinity for the convex face of highly curved regions of synthetic bilayer vesicles, as monitored by circular dichroism spectroscopy [8].

Similarly, a G protein-coupled serotonin receptor (5-HT1A) was found to function more rapidly in a more highly curved membrane [7]. This curvature effect could arise from a better match in the thickness of the hydrophobic portion of the membrane surrounding the protein, stabilizing the active protein form.

In addition to responding to changes in local curvature, membrane proteins can also influence the local curvature of the membrane. Some membrane proteins accomplish this by trafficking individual lipids across the membrane, resulting in uneven lipid compositions across the two membrane leaflets. The membrane must curve so that the leaflet with fewer lipids is on the interior of the curve in order to maintain optimal packing of the head groups [6]. Similarly, membrane leaflets containing lipids with different head group areas may also have some intrinsic curvature due to their differing surface areas [6]. In both cases, the integrity of the membrane as a barrier is facilitated by curvature, protecting the hydrophobic fatty acid tails from the aqueous environment and maintaining separation between the membrane bound compartment and its exterior.

Yet another example of the interplay between membrane proteins and membrane curvature is the process of vesicle formation. The ability to form vesicles is important in the trafficking of newly-formed proteins, the recycling of cell membrane components, and the intake of particles via endocytosis, among countless other cell functions. Vesicle formation involves complex regions of curvature, including low positive curvature when the vesicle is first beginning to form, greater positive curvature as the vesicle swells, and very high negative curvature where the vesicle meets the parent membrane. This complex curvature is facilitated in part by the membrane protein dynamin, which is responsible for creating the &ldquoneck&rdquo separating the parent membrane from the nearly fully-formed vesicle. This is a result of dynamin binding the membrane and taking on a helical conformation, physically forcing the parent membrane to assume a tube-like shape [14]. Coatomers are small proteins that may also play a role in vesicle formation by binding existing membrane proteins in a relatively confined area. The coatomers result in steric repulsion, leading to membrane curvature and budding [6].

Figure (PageIndex<4>): Proteins containing curvature-sensing domains (pink) can play a significant role in vesicle formation, crucial to intercellular and intracellular communication [25].

The binding of amphipathic helices within the membrane is another common mechanism of protein-induced membrane curvature. As shown in the Figure below, one portion of the helix is hydrophobic, while the other is hydrophilic. The hydrophobic portion associates with the fatty acyl chains of the membrane, while the hydrophilic portion interacts with the head groups. This changes the area per head group for the bound leaflet of the membrane, and the membrane must curve in order to compensate.

Figure (PageIndex<5>): Membrane curvature as the result of insertion of an amphipathic helix into one leaflet.

Some of the other well understood ways that lipid-protein interaction relates to curvature of the membrane are discussed in membrane curvature in detail .

Effect of Changes in Membrane Thickness

An important feature of lipids that can influence protein function is the thickness of the bilayer. As shown in Figure (PageIndex<2>), the hydrophobic thickness of a membrane is the distance between the hydrophilic headgroups of either leaflet, which is related to the length of the fatty acid tails of the lipids composing the membrane. Membrane proteins need to match the hydrophobic thickness of the acyl chains for two reasons. First, acyl chains and hydrophobic groups of the membrane proteins do not form hydrogen bonds with water, and tend to minimize their contact with it as this is more thermodynamically favorable (see hydrophobic effect). When the hydrophobic thickness of the membrane and the membrane protein do not match, hydrophobic mismatch occurs. In order to minimize the exposure of hydrophobic portions of the lipid bilayer and transmembrane proteins to the aqueous environment, the bilayer can distort in various ways. It can stretch (Figure (PageIndex<5>)), compress, or even tilt. Stretching involves an extension of the fatty acid chains of the lipids immediately surrounding the protein, providing a greater hydrophobic thickness to better match the size of the protein's hydrophobic domain. This minimizes the extent of hydrophobic membrane protein exposed to the aqueous environment, which is more energetically favorable. Conversely, compression involves a contraction of the fatty acid tails surrounding the protein.

Figure (PageIndex<6>). Stretching of a lipid bilayer to match the hydrophobic thickness of the protein (green). The extension of the fatty acid tails results in a greater hydrophobic thickness, protecting hydrophobic portions of the protein from the hydrophilic environment.

Furthermore, the membrane proteins can aggregate (Figure (PageIndex<3>)) to minimize the area exposed to water [16]. In addition to being more thermodynamically favorable (i.e. maximizing entropy by reducing the number of water molecules in highly ordered contact around hydrophobic residues), avoiding exposure of a transmembrane protein to aqueous environment is also important for optimal function because the hydrophobic side chains of the protein can change conformation when they contact water, distorting their native structure, and causing functional loss. The studies on these membrane distortions carried out with model membranes (artificial bilayers), show that the thickness of the acyl chain region is an essential part of protein function. For instance, sarcoplasmic reticulum Ca-ATPase, which is a protein important in calcium regulation in muscles, was shown to be greatly affected by the number of the carbons in the acyl chain, which determines the hydrophobic thickness of the membrane [16]. In addition, molecular dynamics simulations showed the possibility of protein aggregation (Figure (PageIndex<3>)) due to hydrophobic mismatch [17].

Figure (PageIndex<7>). Aggregated proteins to minimize hydrophobic mismatch [21].

Membrane proteins may also change conformation in response to changes in membrane tension, which can be indicative of changing external conditions necessitating a response from the cell. For example, osmotic swelling or shrinking of a cell will result in higher or lower membrane tension, respectively. Under increased tension, the bilayer will tend to thin, resulting in increased hydrophobic mismatch. One possibility to mitigate this is for the membrane conformation to change so that its hydrophobic domain thickness matches that of the transmembrane protein. Another possibility is that a conformational change in the membrane protein structure could relieve hydrophobic mismatch, making a response to membrane tension an effective gating mechanism. This is typically more energetically favorable in the case of significant membrane deformations, for which the energy required to change the membrane conformation to match the protein&rsquos hydrophobic thickness would be greater than the cost of changing the protein&rsquos conformation [11]. Such conformational changes include changes in the angle at which the transmembrane domain is tilted or unwinding of alpha-helical transmembrane domains [11]. The response of membrane proteins to hydrophobic mismatch arising from membrane tension is evident in the case of many bacterial mechanosensitive channels, which under higher membrane tension are more likely to be open [15].

For example, the lipid-protein interaction energy for a membrane protein with six helical transmembrane domains has been estimated to increase by approximately 10 -20 J/lipid molecule for a 10 Å hydrophobic mismatch, providing an appreciable driving force for either the membrane or protein to change its conformation [11]. At room temperature, the energetic cost of bending a lipid membrane is typically on the order of 10 -20 J, although this can change significantly based on the local temperature and lipid composition [24]. Hydrolyzing one ATP, a typical means of driving protein conformational changes, yields approximately 10 -19 J [6]. All these values are within similar orders of magnitude, suggesting the most effective method of resolving hydrophobic mismatch for a given membrane and protein may depend on both intrinsic and environmental factors.

The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Abankwa, D., Hanzal-Bayer, M., Ariotti, N., Plowman, S. J., Gorfe, A. A., Parton, R. G., et al. (2008). A novel switch region regulates H-ras membrane orientation and signal output. EMBO J. 27, 727�. doi: 10.1038/emboj.2008.10

Abdullah, U., and Cullen, P. J. (2009). The tRNA modification complex elongator regulates the Cdc42-dependent mitogen-activated protein kinase pathway that controls filamentous growth in yeast. Eukaryot. Cell 8, 1362�. doi: 10.1128/EC.00015-09

Aittoniemi, J., Róg, T., Niemelä. P, Pasenkiewicz-Gierula, M., Karttunen, M., and Vattulainen, I. (2006). Tilt: major factor in sterols' ordering capability in membranes. J. Phys. Chem. B 110, 25562�. doi: 10.1021/jp064931u

Alexander, R. T., and Grinstein, S. (2006). Na+/H+ exchangers and the regulation of volume. Acta Physiol. 187, 159�. doi: 10.1111/j.1748-1716.2006.01558.x

Andersen, O. S., and Koeppe, R. E. II. (2007). Bilayer thickness and membrane protein function: an energetic perspective. Annu. Rev. Biophys. Biomol. Struct. 36, 107�. doi: 10.1146/annurev.biophys.36.040306.132643

Anishkin, A., Loukin, S. H., Teng, J., and Kung, C. (2014). Feeling the hidden mechanical forces in lipid bilayer is an original sense. Proc. Natl. Acad. Sci. U.S.A. 111, 7898�. doi: 10.1073/pnas.1313364111

Bader, M. W., Sanowar, S., Daley, M. E., Schneider, A. R., Cho, U., Xu, W., et al. (2005). Recognition of antimicrobial peptides by a bacterial sensor kinase. Cell 122, 461�. doi: 10.1016/j.cell.2005.05.030

Ballou, E. R., Kozubowski, L., Nichols, C. B., and Alspaugh, J. A. (2013) Ras1 acts through duplicated Cdc42 Rac proteins to regulate morphogenesis pathogenesis in the human fungal pathogen Cryptococcus neoformans. PLoS Genet. 9:e1003687. doi: 10.1371/journal.pgen.1003687

Basu, R., Munteanu, E. L., and Chang, F. (2014). Role of turgor pressure in endocytosis in fission yeast. Mol. Biol. Cell 25, 679�. doi: 10.1091/mbc.e13-10-0618

Battle, A. R., Ridone, P., Bavi, N., Nakayama, Y., Nikolaev, Y. A., and Martinac, B. (2015). Lipid-protein interactions: lessons learned from stress. Biochim. Biophys. Acta 1848, 1744�. doi: 10.1016/j.bbamem.2015.04.012

Bavi, N., Cox, C. D., Perozo, E., and Martinac, B. (2017). Toward a structural blueprint for bilayer-mediated channel mechanosensitivity. Channels 11, 91�. doi: 10.1080/19336950.2016.1224624

Belenky, P., Camacho, D., and Collins, J. J. (2013). Fungicidal drugs induce a common oxidative-damage cellular death pathway. Cell Rep. 3, 350�. doi: 10.1016/j.celrep.2012.12.021

Belotti, F., Tisi, R., and Martegani, E. (2006). The N-terminal region of the Saccharomyces cerevisiae RasGEF Cdc25 is required for nutrient-dependent cell-size regulation. Microbiology 152, 1231�. doi: 10.1099/mic.0.28683-0

Belotti, F., Tisi, R., Paiardi, C., Rigamonti, M., Groppi, S., and Martegani, E. (2012). Localization of Ras signaling complex in budding yeast. Biochim. Biophys. Acta 1823, 1208�. doi: 10.1016/j.bbamcr.2012.04.01

Bhate, M. P., Lemmin, T., Kuenze, G., Mensa, B., Ganguly, S., Peters, J. M., et al. (2018). Structure and function of the transmembrane domain of NsaS, an antibiotic sensing histidine kinase in Staphylococcus aureus. J. Am. Chem. Soc. 140, 7471�. doi: 10.1021/jacs.7b09670

Bhate, M. P., Molnar, K. S., Goulian, M., and DeGrado, W. F. (2015). Signal transduction in histidine kinases: insights from new structures. Structure 23, 981�. doi: 10.1016/j.str.2015.04.002

Bond, M., Croft, W., Tyson, R., Bretschneider, T., Davey, J., and Ladds, G. (2013). Quantitative analysis of human ras localization and function in the fission yeast Schizosaccharomyces pombe. Yeast 30, 145�. doi: 10.1002/yea.2949

Brewster, J. L., and Gustin, M. C. (2014). Hog1: 20 years of discovery and impact. Sci. Signal. 7:re7. doi: 10.1126/scisignal.2005458

Broggi, S., Martegani, E., and Colombo, S. (2013). Nuclear Ras2-GTP controls invasive growth in Saccharomyces cerevisiae. PLoS ONE 8:e79274. doi: 10.1371/journal.pone.0079274

Busti, S., Coccetti, P., Alberghina, L., and Vanoni, M. (2010). Glucose signaling-mediated coordination of cell growth and cell cycle in Saccharomyces cerevisiae. Sensors 10, 6195�. doi: 10.3390/s100606195

Cannatelli, A., Giani, T., Dɺndrea, M. M., Di Pilato, V., Arena, F., Conte, V., et al. (2014). MgrB inactivation is a common mechanism of colistin resistance in KPC-producing Klebsiella pneumoniae of clinical origin. Antimicrob. Agents Chemother. 58, 5696�. doi: 10.1128/AAC.03110-14

Chang, F. (2017). Forces that shape fission yeast cells. Mol. Biol. Cell 28, 1819�. doi: 10.1091/mbc.e16-09-0671

Chen, R. E., and Thorner, J. (2007). Function and regulation in MAPK signaling pathways:lessons learned from the yeast Saccharomyces cerevisiae. Biochim. Biophys. Acta 1773, 1311�. doi: 10.1016/j.bbamcr.2007.05.003

Cho, U. S., Bader, M. W., Amaya, M. F., Daley, M. E., Klevit, R. E., Miller, S. I., et al. (2006). Metal bridges between the PhoQ sensor domain and the membrane regulate transmembrane signaling. J. Mol. Biol. 356, 1193�. doi: 10.1016/j.jmb.2005.12.032

Christensen, S. M., Tu, H. L., Jun, J. E., Alvarez, S., Triplet, M. G., Iwig, J. S., et al. (2016). One-way membrane trafficking of SOS in receptor-triggered Ras activation. Nat. Struct. Mol. Biol. 23, 838�. doi: 10.1038/nsmb.3275

Clifton, L. A., Skoda, M. W., Le Brun, A. P., Ciesielski, F., Kuzmenko, I., Holt, S. A., et al. (2015). Effect of divalent cation removal on the structure of gram-negative bacterial outer membrane models. Langmuir 31, 404�. doi: 10.1021/la504407v

Cohen, B. E. (2016). The role of signaling via aqueous pore formation in resistance responses to amphotericin B. Antimicrob. Agents Chemother. 60, 5122�. doi: 10.1128/AAC.00878-16

Copp, J., Wiley, S., Ward, M. W., and van der Geer, P. (2005). Hypertonic shock inhibits growth factor receptor signaling, induces caspase-3 activation, and causes reversible fragmentation of the mitochondrial network. Am. J. Physiol. Cell Physiol. 288, C403�. doi: 10.1152/ajpcell.00095.2004

Cullen, P. J., and Sprague, G. F. Jr. (2012). The regulation of filamentous growth in yeast. Genetics 190, 23�. doi: 10.1534/genetics.111.127456

Davenport, K. D., Williams, K. E., Ullmann, B. D., and Gustin, M. C. (1999). Activation of the Saccharomyces cerevisiae filamentation/invasion pathway by osmotic stress in high-osmolarity glycogen pathway mutants. Genetics 153, 1091�.

DeFeo-Jones, D., Tatchell, K., Robinson, L. C., Sigal, I. S., Vass, W. C., Lowy, D. R., et al. (1985). Mammalian and yeast ras gene products: biological function in their heterologous systems. Science 228, 179�.

Dickson, R. C., Sumanasekera, C., and Lester, R. L. (2006). Functions and metabolism of sphingolipids in Saccharomyces cerevisiae. Prog. Lipid Res. 45, 447�. doi: 10.1016/j.plipres.2006.03.004

Eisenberg, S., and Henis, Y. I. (2008). Interactions of Ras proteins with the plasma membrane and their roles in signaling. Cell. Signal. 20, 31�. doi: 10.1016/j.cellsig.2007.07.012

Elmore, D. E., and Dougherty, D. A. (2003). Investigating lipid composition effects on the mechanosensitive channel of large conductance (MscL) using molecular dynamics simulations. Biophys. J. 85, 1512�. doi: 10.1016/S0006-3495(03)74584-6

Falke, J. J. (2014). Piston versus scissors: chemotaxis receptors versus sensor His-kinase receptors in two-component signaling pathways. Structure 22, 219�. doi: 10.1016/j.str.2014.08.011

Gao, R., and Stock, A. M. (2009). Biological insights from structures of two-component proteins. Annu. Rev. Microbiol. 63, 133�. doi: 10.1146/annurev.micro.091208.073214

Garc໚-Sพz, A. J., Chiantia, S., and Schwille, P. (2007). Effect of line tension on the lateral organization of lipid membranes. J. Biol. Chem. 282, 33537�. doi: 10.1074/jbc.M706162200

Gimeno, C. J., Ljungdahl, P. O., Styles, C. A., and Fink, G. R. (1992). Unipolar cell divisions in the yeast S. cerevisiae lead to filamentous growth: regulation by starvation and RAS. Cell 68, 1077�. doi: 10.1016/0092-8674(92)90079-R

Gorfe, A. A., Babakhani, A., and McCammon, J. A. (2007a). H-ras protein in a bilayer: interaction and structure perturbation. J. Am. Chem. Soc. 129, 12280�. doi: 10.1021/ja073949v

Gorfe, A. A., Hanzal-Bayer, M., Abankwa, D., Hancock, J. F., and McCammon, J. A. (2007b). Structure and dynamics of the full-length lipid-modified H-Ras protein in a 1,2-dimyristoylglycero-3-phosphocholine bilayer. J. Med. Chem. 50, 674�. doi: 10.1021/jm061053f

Grage, S. L., Afonin, S., Kara, S., Buth, G., and Ulrich, A. S. (2016). Membrane thinning and thickening induced by membrane-active amphipathic peptides. Front. Cell Dev. Biol. 4:65. doi: 10.3389/fcell.2016.00065

Grecco, H. E., Schmick, M., and Bastiaens, P. I. (2011). Signaling from the living plasma membrane. Cell 144, 897�. doi: 10.1016/j.cell.2011.01.029

Groves, J. T., and Kuriyan, J. (2010). Molecular mechanisms in signal transduction at the membrane. Nat. Struct. Mol. Biol. 17, 659�. doi: 10.1038/nsmb.1844

Gureasko, J., Galush, W. J., Boykevisch, S., Sondermann, H., Bar-Sagi, D., Groves, J. T., et al. (2008). Membrane-dependent signal integration by the Ras activator Son of sevenless. Nat. Struct. Mol. Biol. 15, 452�. doi: 10.1038/nsmb.1418

Gushchin, I., and Gordeliy, V. (2018). Transmembrane signal transduction in two-component systems: piston, scissoring, or helical rotation? Bioessays 40, 1�. doi: 10.1002/bies.201700197

Gutin, J., Sadeh, A., Rahat, A., Aharoni, A., and Friedman, N. (2015). Condition-specific genetic interaction maps reveal crosstalk between the cAMP/PKA and the HOG MAPK pathways in the activation of the general stress response. Mol. Syst. Biol. 11:829. doi: 10.15252/msb.20156451

Guzmán, C., Šolman, M., Ligabue, A., Bla៞vitš, O., Andrade, D. M., Reymond, L., et al. (2014). The efficacy of Raf kinase recruitment to the GTPase H-ras depends on H-ras membrane conformer-specific nanoclustering. J. Biol. Chem. 289, 9519�. doi: 10.1074/jbc.M113.537001

Hancock, J. F. (2003). Ras proteins: different signals from different locations. Nat. Rev. Mol. Cell Biol. 4, 73�. doi: 10.1038/nrm1105

Hancock, J. F. (2006) Lipid rafts: contentious only from simplistic standpoints. Nat. Rev. Mol. Cell Biol. 7, 456�. doi: 10.1038/nrm1925

Hardie, R. C., and Franze, K. (2012). Photomechanical responses in Drosophila photoreceptors. Science 338, 260�. doi: 10.1126/science.1222376

Henis, Y. I., Hancock, J. F., and Prior, I. A. (2009). Ras acylation, compartmentalization and signaling nanoclusters. Mol. Membr. Biol. 26, 80�. doi: 10.1080/09687680802649582

Henzler-Wildman, K. A., Martinez, G. V., Brown, M. F., and Ramamoorthy, A. (2004). Perturbation of the hydrophobic core of lipid bilayers by the human antimicrobial peptide LL-37. Biochemistry 43, 8459�. doi: 10.1021/bi036284s

Hooley, R., Yu, C. Y., Symons, M., and Barber, D. L. (1996). G alpha 13 stimulates Na + -H + exchange through distinct Cdc42-dependent and RhoA-dependent pathways. J. Biol. Chem. 271, 6152�. doi: 10.1074/jbc.271.11.6152

Horie, T., Tatebayashi, K., Yamada, R., and Saito, H. (2008). Phosphorylated Ssk1 prevents unphosphorylated Ssk1 from activating the Ssk2 mitogen-activated protein kinase kinase kinase in the yeast high-osmolarity glycerol osmoregulatory pathway. Mol. Cell. Biol. 17, 5172�. doi: 10.1128/MCB.00589-08

Huang, S., and Ingber, D. E. (2005). Cell tension, matrix mechanics, and cancer Development 8, 175�. doi: 10.1016/j.ccr.2005.08.009

Jang, H., Banerjee, A., Chavan, T. S., Lu, S., Zhang, J., Gaponenko, V., et al. (2016). The higher level of complexity of K-Ras4B activation at the membrane. FASEB J. 30, 1643�. doi: 10.1096/fj.15-279091

Kamada, Y., Jung, U. S., Piotrowski, J., and Levin, D. E. (1995). The protein kinase C-activated MAP kinase pathway of Saccharomyces cerevisiae mediates a novel aspect of the heat shock response. Genes Dev. 9, 1559�. doi: 10.1101/gad.9.13.1559

Kapoor, S., Werkmüller, A., Goody, R. S., Waldmann, H., and Winter, R. (2013). Pressure modulation of Ras-membrane interactions and intervesicle transfer. J. Am. Chem. Soc. 135, 6149�. doi: 10.1021/ja312671j

Killian, J. A., and von Heijne, G. (2000). How proteins adapt to a membrane-water interface. Trends Biochem. Sci. 25, 429�. doi: 10.1016/S0968-0004(00)01626-1

Klose, C., Ejsing, C. S., Garc໚-Sพz, A. J., Kaiser, H. J., Sampaio, J. L., Surma, M. A., et al. (2010). Yeast lipids can phase-separate into micrometer-scale membrane domains. J. Biol. Chem. 285, 30224�. doi: 10.1074/jbc.M110.123554

Kock, C., Arlt, H., Ungermann, C., and Heinisch, J. J. (2016). Yeast cell wall integrity sensors form specific plasma membrane microdomains important for signaling. Cell. Microbiol. 18, 1251�. doi: 10.1111/cmi.12635

Langosch, D., and Arkin, I. T. (2009). Interaction and conformational dynamics of membrane-spanning protein helices. Protein Sci. 18, 1343�. doi: 10.1002/pro.154

Larsen, J. B., Jensen, M. B., Bhatia, V. K., Pedersen, S. L., Bjørnholm, T., Iversen, L., et al. (2015). Membrane curvature enables N-Ras lipid anchor sorting to liquid-ordered membrane phases. Nat. Chem. Biol. 11, 192�. doi: 10.1038/nchembio.1733

Levin, D. E. (2005). Cell wall integrity signaling in Saccharomyces cerevisiae. Microbiol. Mol. Biol. Rev. 69, 262�. doi: 10.1128/MMBR.69.2.262-291.2005

Li, S., Zhang, X., and Wang, W. (2010). Cluster formation of anchored proteins induced by membrane-mediated interaction. Biophys. J. 98, 2554�. doi: 10.1016/j.bpj.2010.02.032

Lin, X., Li, Z., and Gorfe, A. A. (2015). Reversible effects of peptide concentration and lipid composition on h-ras lipid anchor clustering. Biophys. J. 109, 2467�. doi: 10.1016/j.bpj.2015.11.009

Linder, M. E., and Deschenes, R. J. (2003). New insights into the mechanisms of protein palmitoylation. Biochemistry 42, 4311�. doi: 10.1021/bi034159a

Lippa, A. M., and Goulian, M. (2009). Feedback inhibition in the PhoQ/PhoP signaling system by a membrane peptide. PLoS Genet. 5:e1000788. doi: 10.1371/journal.pgen.1000788.

Liu, W. F., Nelson, C. M., Tan, J. L., and Chen, C. S. (2007). Cadherins, RhoA, and Rac1 are differentially required for stretch-mediated proliferation in endothelial versus smooth muscle cells. Circ. Res. 101:e44�. doi: 10.1161/CIRCRESAHA.107.158329

Lorenz, M. C., Pan, X., Harashima, T., Cardenas, M. E., Xue, Y., Hirsch, J. P., et al. (2010). The G protein-coupled receptor gpr1 is a nutrient sensor that regulates pseudohyphal differentiation in Saccharomyces cerevisiae. Genetics 154, 609�.

Lu, S., Jang, H., Muratcioglu, S., Gursoy, A., Keskin, O., Nussinov, R., et al (2016). Ras conformational ensembles, allostery, and signaling. Chem. Rev. 116, 6607�. doi: 10.1021/acs.chemrev.5b00542

Lundbaek, J. A., Collingwood, S. A., Ingólfsson, H. I., Kapoor, R., and Andersen, O. S. (2010). Lipid bilayer regulation of membrane protein function: gramicidin channels as molecular force probes. J. R. Soc. Interface 7, 373�. doi: 10.1098/rsif.2009.0443

Maeng, S., Ko, Y. J., Kim, G. B., Jung, K. W., Floyd, A., Heitman, J., et al. (2010). Comparative transcriptome analysis reveals novel roles of the Ras and cyclic AMP signaling pathways in environmental stress response and antifungal drug sensitivity in Cryptococcus neoformans. Eukaryotic Cell 9, 360�. doi: 10.1128/EC.00309-09

Martinac, B., Adler, J., and Kung, C. (1990). Mechanosensitive ion channels of E. coli activated by amphipaths. Nature 348, 261�. doi: 10.1038/348261a0

Mascher, T., Helmann, J. D., and Unden, G. (2006). Stimulus perception in bacterial signal-transducing histidine kinases. Microbiol. Mol. Biol. Rev. 70, 910�. doi: 10.1128/MMBR.00020-06

Mecke, A., Lee, D. K., Ramamoorthy, A., Orr, B. G., and Banaszak Holl, M. M. (2005). Membrane thinning due to antimicrobial peptide binding: an atomic force microscopy study of MSI-78 in lipid bilayers. Biophys. J. 89, 4043�. doi: 10.1529/biophysj.105.062596

Merchan, S., Berna, D., Serrano, R., and Yenush, L. (2004). Response of the Saccharomyces cerevisiae Mpk1 mitogen-activated protein kinase pathway to increases in internal turgor pressure caused by loss of Ppz protein phosphatases. Eukaryotic Cell 3, 100�. doi: 10.1128/EC.3.1.100-107.2004

Mollinedo, F. (2012). Lipid raft involvement in yeast cell growth and death. Front. Oncol. 2:140. doi: 10.3389/fonc.2012.00140

Molnar, K. S., Bonomi, M., Pellarin, R., Clinthorne, G. D., Gonzalez, G., Goldberg, S. D., et al. (2014). Cys-scanning disulfide crosslinking and Bayesian modeling probe the transmembrane signaling mechanism of the histidine kinase, PhoQ. Structure 22, 1239�. doi: 10.1016/j.str.2014.04.019

Nichols, C. B., Ferreyra, J., Ballou, E. R., and Alspaugh, J. A. (2009). Subcellular localization directs signaling specificity of the Cryptococcus neoformans Ras1 protein. Eukaryotic Cell 8, 181�. doi: 10.1128/EC.00351-08

Nichols, C. B., Perfect, Z. H., and Alspaugh, J. A. (2007). A Ras1-Cdc24 signal transduction pathway mediates thermotolerance in the fungal pathogen Cryptococcus neoformans. Mol. Microbiol. 63, 1118�. doi: 10.1111/j.1365-2958.2006.05566.x

Norimatsu, Y., Hasegawa, K., Shimizu, N., and Toyoshima, C. (2017). Protein-phospholipid interplay revealed with crystals of a calcium pump. Nature 545, 193�. doi: 10.1038/nature22357

Nussinov, R., Tsai, C. J., and Jang, H. (2018). Oncogenic Ras isoforms signaling specificity at the membrane. Cancer Res. 78, 593�. doi: 10.1158/0008-5472.CAN-17-2727

Onken, B., Wiener, H., Philips, M. R., and Chang, E. C. (2006). Compartmentalized signaling of Ras in fission yeast. Proc. Natl. Acad. Sci. U.S.A. 103, 9045�. doi: 10.1073/pnas.0603318103

Ozdemir, E. S., Jang, H., Gursoy, A., Keskin, O., Li, Z., and Sacks, D. B. (2018). Unraveling the molecular mechanism of interactions of the Rho GTPases Cdc42 and Rac1 with the scaffolding protein IQGAP2. J. Biol. Chem. 293, 3685�. doi: 10.1074/jbc.RA117.001596

Papaleo, E. (2015). Integrating atomistic molecular dynamics simulations, experiments, and network analysis to study protein dynamics: strength in unity. Front. Mol. Biosci. 27:28. doi: 10.3389/fmolb.2015.00028

Perozo, E., Cortes, D. M., Sompornpisut, P., Kloda, A., and Martinac, B. (2002). Open channel structure of MscL and the gating mechanism of mechanosensitive channels. Nature 418, 942�. doi: 10.1038/nature00992

Phillips, M. J., Calero, G., Chan, B., Ramachandran, S., and Cerione, R. A. (2008). Effector proteins exert an important influence on the signaling-active state of the small GTPase Cdc42. J. Biol. Chem. 283, 14153�. doi: 10.1074/jbc.M706271200

Prior, I. A., Harding, A., Yan, J., Sluimer, J., Parton, R. G., and Hancock, J. F. (2001). GTP-dependent segregation of H-ras from lipid rafts is required for biological activity. Nat. Cell Biol. 3, 368�. doi: 10.1038/35070050

Reiser, V., Raitt, D. C., and Saito, H. (2003). Yeast osmosensor Sln1 and plant cytokinin receptor Cre1 respond to changes in turgor pressure. J. Cell Biol. 161, 1035�. doi: 10.1083/jcb.200301099

Remorino, A., De Beco, S., Cayrac, F., Di Federico, F., Cornilleau, G., Gautreau, A., et al. (2017). Gradients of Rac1 nanoclusters support spatial patterns of Rac1 signaling. Cell Rep. 21, 1922�. doi: 10.1016/j.celrep.2017.10.069

Rosenberger, C. M., Gallo, R. L., and Finlay, B. B. (2004). Interplay between antibacterial effectors: a macrophage antimicrobial peptide impairs intracellular Salmonella replication. Proc. Natl. Acad. Sci. U.S.A. 101, 2422�. doi: 10.1073/pnas.0304455101

Rotblat, B., Prior, I. A., Muncke, C., Parton, R. G., Kloog, Y., Henis, Y. I., et al. (2004). Three separable domains regulate GTP-dependent association of H-ras with the plasma membrane. Mol. Cell. Biol. 24, 6799�. doi: 10.1128/MCB.24.15.6799-6810.2004

Ruckwardt, T., Scott, A., Scott, J., Mikulecky, P., and Hartsel, S. C. (1998). Lipid and stress dependence of amphotericin B ion selective channels in sterol-free membranes. Biochim. Biophys. Acta 1372, 283�. doi: 10.1016/S0005-2736(98)00073-X

Rudoni, S., Colombo, S., Coccetti, P., and Martegani, E. (2001). Role of guanine nucleotides in the regulation of the Ras/cAMP pathway in Saccharomyces cerevisiae. Biochim. Biophys. Acta 1538, 181�. doi: 10.1016/S0167-4889(01)00067-2

Saito, H., and Posas, F. (2012). Response to hyperosmotic stress. Genetics 192, 289�. doi: 10.1534/genetics.112.140863

Schaber, J., Adrover, M. A., Eriksson, E., Pelet, S., Petelenz-Kurdziel, E., Klein, D., et al. (2010). Biophysical properties of Saccharomyces cerevisiae and their relationship with HOG pathway activation. Eur. Biophys. J. 39, 1547�. doi: 10.1007/s00249-010-0612-0

Schliess, F., Reinehr, R., and Häussinger, D. (2007). Osmosensing and signaling in the regulation of mammalian cell function. FEBS J. 274, 5799�. doi: 10.1111/j.1742-4658.2007.06100.x

Schmidt, D., del Mármol, J., and MacKinnon, R. (2012). Mechanistic basis for low threshold mechanosensitivity in voltage-dependent K+ channels. Proc. Natl. Acad. Sci. U.S.A. 109, 10352�. doi: 10.1073/pnas.1204700109

Schneiter, R., Brügger, B., Amann, C. M., Prestwich, G. D., Epand, R. F., Zellnig, G., et al. (2004). Identification and biophysical characterization of a very-long-chain-fatty-acid-substituted phosphatidylinositol in yeast subcellular membranes. Biochem. J. 381, 941�. doi: 10.1042/BJ20040320

Shatursky, O. Y., Romanenko, O. V., and Himmelreich, N. H. (2014). Long open amphotericin channels revealed in cholesterol-containing phospholipid membranes are blocked by thiazole derivative. J. Membr. Biol. 247, 211�. doi: 10.1007/s00232-013-9626-8

Simi, A. K., Piotrowski, A. S., and Nelson, C. M. (2015). “Mechanotransduction, Metastasis and Genomic Instability,” in Genomic Instability and Cancer Metastasis, eds C. Maxwell and C. Roskelley (Switzerland: Springer International Publishing), 139�.

Simons, K., and Sampaio, J. L. (2011). Membrane organization and lipid rafts. Cold Spring Harb. Perspect. Biol. 3:a004697. doi: 10.1101/cshperspect.a004697

Son, S., Kang, J. H., Oh, S., Kirschner, M. W., Mitchison, T. J., and Manalis, S. (2015). Resonant microchannel volume and mass measurements show that suspended cells swell during mitosis. J. Cell Biol. 211, 757�. doi: 10.1083/jcb.201505058

Straede, A., Corran, A., Bundy, J., and Heinisch, J. J. (2007). The effect of tea tree oil and antifungal agents on a reporter for yeast cell integrity signaling. Yeast 24, 321�. doi: 10.1002/yea.1478

Sukharev, S. I., Blount, P., Martinac, B., and Kung, C. (1997). Mechanosensitive channels of Escherichia coli: the MscL gene, protein, and activities. Annu. Rev. Physiol. 59, 633�. doi: 10.1146/annurev.physiol.59.1.633

Tamanoi, F. (2011). Ras signaling in yeast. Genes Cancer 2, 210�. doi: 10.1177/1947601911407322

Tanigawa, M., Kihara, A., Terashima, M., Takahara, T., and Maeda, T. (2012). Sphingolipids regulate the yeast high-osmolarity glycerol response pathway. Mol. Cell. Biol. 32, 2861�. doi: 10.1128/MCB.06111-11

Tatebayashi, K., Tanaka, K., Yang, H. Y., Yamamoto, K., Matsushita, Y., Tomida, T., et al. (2007). Transmembrane mucins Hkr1 and Msb2 are putative osmosensors in the SHO1 branch of yeast HOG pathway. EMBO J. 26, 3521�. doi: 10.1038/sj.emboj.7601796

Tatebayashi, K., Yamamoto, K., Nagoya, M., Takayama, T., Nishimura, A., Sakurai, M., et al. (2015). Osmosensing and scaffolding functions of the oligomeric four-transmembrane domain osmosensor Sho1. Nat. Commun. 21:6975. doi: 10.1038/ncomms7975

Teixeira-Santos, R., Ricardo, E., Branco, R. J., Azevedo, M. M., Rodrigues, A. G., and Pina-Vaz, C. (2016). Unveiling the synergistic interaction between liposomal amphotericin B and colistin. Front. Microbiol. 7:1439. doi: 10.3389/fmicb.2016.01439

Te Welscher, Y. M., ten Napel, H. H., Balagué, M. M., Souza, C. M., Riezman, H., de Kruijff, B., et al. (2008). Natamycin blocks fungal growth by binding specifically to ergosterol without permeabilizing the membrane. J. Biol. Chem. 283, 6393�. doi: 10.1074/jbc.M707821200

Toledo, A., Huang, Z., Coleman, J. L., London, E., and Benach, J. L. (2018). Lipid rafts can form in the inner and outer membranes of Borrelia burgdorferi and have different properties and associated proteins. Mol. Microbiol. 108, 63�. doi: 10.1111/mmi.13914

Vadaie, N., Dionne, H., Akajagbor, D. S., Nickerson, S. R., Krysan, D. J., and Cullen, P. J. (2008). Cleavage of the signaling mucin Msb2 by the aspartyl protease Yps1 is required for MAPK activation in yeast. J. Cell Biol. 181, 1073�. doi: 10.1083/jcb.200704079

Van der Wijk, T., Dorrestijn, J., Narumiya, S., Maassen, J. A., de Jonge, H. R., and Tilly, B. C. (1998). Osmotic swelling-induced activation of the extracellular-signal-regulated protein kinases Erk-1 and Erk-2 in intestine 407 cells involves the Ras/Raf-signaling pathway. Biochem. J. 331, 863�. doi: 10.1042/bj3310863

Verna, J., Lodder, A., Lee, K., Vagts, A., and Ballester, R. (1997). A family of genes required for maintenance of cell wall integrity and for the stress response in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. U.S.A. 94, 13804�.

Waugh, M. S., Nichols, C. B., DeCesare, C. M., Cox, G. M., Heitman, J., and Alspaugh, J. A. (2002). Ras1 and Ras2 contribute shared and unique roles in physiology and virulence of Cryptococcus neoformans. Microbiology 148, 191�. doi: 10.1099/00221287-148-1-191

Weise, K., Triola, G., Janosch, S., Waldmann, H., and Winter, R. (2010). Visualizing association of lipidated signaling proteins in heterogeneous membranes–partitioning into subdomains, lipid sorting, interfacial adsorption, and protein association. Biochim. Biophys. Acta 1798, 1409�. doi: 10.1016/j.bbamem.2009.12.006

West, A. H., and Stock, A. M. (2001). Histidine kinases and response regulator proteins in two-component signaling systems. Trends Biochem. Sci. 26, 369�. doi: 10.1016/S0968-0004(01)01852-7

Yadavalli, S. S., Carey, J. N., Leibman, R. S., Chen, A. I., Stern, A. M., Roggiani, M., et al. (2016). Antimicrobial peptides trigger a division block in Escherichia coli through stimulation of a signaling system. Nat. Commun. 29:12340. doi: 10.1038/ncomms12340

Yang, H. Y., Tatebayashi, K., Yamamoto, K., and Saito, H. (2009). Glycosylation defects activate filamentous growth Kss1, MAPK and inhibit osmoregulatory Hog1 MAPK. EMBO J. 28, 1380�. doi: 10.1038/emboj.2009.104

Yusuf, R., Nguyen, T. L., Heininger, A., Lawrence, R. J., Hall, B. A., and Draheim, R. R. (2017). In vivo cross-linking and transmembrane helix dynamics support a bidirectional non-piston model of signaling within E. coli EnvZ. bioRxiv 206888. doi: 10.1101/206888

Zhou, Y., and Hancock, J. F. (2015). Ras nanoclusters: versatile lipid-based signaling platforms. Biochim. Biophys. Acta 1853, 841�. doi: 10.1016/j.bbamcr.2014.09.008

Zlotek-Zlotkiewicz, E., Monnier, S., Cappello, G., Le Berre, M., and Piel, M. (2015). Optical volume and mass measurements show that mammalian cells swell during mitosis. J. Cell Biol. 211, 765�. doi: 10.1083/jcb.201505056

Keywords: osmosensors, lipid bilayer thickness, membrane thinning, Ras proteins, signaling pathways, antimicrobial peptides, colistin, amphotericin B

Citation: Cohen BE (2018) Membrane Thickness as a Key Factor Contributing to the Activation of Osmosensors and Essential Ras Signaling Pathways. Front. Cell Dev. Biol. 6:76. doi: 10.3389/fcell.2018.00076

Received: 27 April 2018 Accepted: 25 June 2018
Published: 24 July 2018.

Manuel Prieto, Universidade de Lisboa, Portugal

Jesus Perez-Gil, Complutense University of Madrid, Spain
Erdinc Sezgin, University of Oxford, United Kingdom

Copyright © 2018 Cohen. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

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