from Montal, Darszon, and Schindler, 1981, Quarterly Reviews of Biophysics 14:1-79
Table of contents
II Determining physical
properties of membrane proteins
5. Isolation, purification, and gel electrophoresis of membrane protein complexes
Critical micellar concentration
Light scattering
Isolation and purification of membrane proteins
Principles of gel electrophoresis
2D Gel electrophoresis
Proteomics
Mass spectrometry
6. Structure determination of membrane proteins
Circular dichroism to study secondary structures of proteins
Studying protein complex assembly - gap junction proteins
GFP - green fluorescence protein
7. Functional membrane protein reconstitution
Membrane vesicles
Planar lipid membranes
Part I Thermodynamics and solution behavior of macromolecules
1. Energy of biological systems
2. Molecular interpretation of thermodynamic quantities
3. Surface tension and phospholipid monolayers
4. Thermodynamic stability of monolayers
Part III Membrane transport
8. Electrodiffusion through ion channels
9. Kinetic models of ion flux across channels
10. Physical models of ion flux across channels
There is a simple experimental need to isolate and purify [membrane] proteins. Biological systems are complex and it is normally difficult to impossible to determine the structure function relationship of a protein in its native environment (in vivo). As a first step, therefore, proteins are reconstituted into simpler systems (in vitro) by removing them from their native surroundings. This allows the determination of intrinsic properties of a protein, properties which are largely independent of other membrane components; molecular weight, charge, secondary structure composition, and often also its function as receptor, pump, transporter, or channel.
Properties of proteins studied in reconstitution systems are called intrinsic because they depend on a minimal set of other molecules; a protein in aqueous solution, or if it is a membrane protein, a detergent extract. The ionic strength, pH, temperature and protein concentration comprise the few variables. An in vitro system thus has clearly defined parameters and the measurement of macroscopic properties such as heat capacity, UV absorption, circular dichroism spectra can easily be interpreted for moclecular structure and mechanisms. Once certain properties have been established as being 'intrinisic' to the protein, its function can be studied in vivo and the influence of other cellular components on its activity determined.
Fig. Outline of reconstitution pathway for ion channels
from Montal, Darszon, and Schindler, 1981, Quarterly
Reviews of Biophysics 14:1-79
Critical micellar concentration
While it is impossible to keep membrane proteins in a polar solvent,
their are many ways to isolate them from their native membrane environment,
purify them to homogeneity (remove all other proteins), and reconstitute
purified proteins into supra molecular structures mimicking either solubility,
or orientation, or both properties of membranes. Removing or isolating
membrane components from native membranes is done with the help of detergents.
Detergents are small amphipathic molecules that have a propensity to form
micelles in aqueous solution. Micelles are small, spherical particles of
a few dozens to hundreds of monomers with a hydrophobic core and a hydrophilic
surface. Detergents are found in three phases all of which are in chemical
equilibrium:
- free monomers
- micelles
- monolayer at air water interface
While the monolayer for most practical purposes is negligible, micellare
structures are determined by studying the phase transition behavior between
monomeric and oligomeric forms. This transition point is characteristic
for each type of detergent and is known as the critial micellar concnetration,
or CMC.
This figure shows typical behvior of three solution properties as a function of detergent concentration. As can be seen, the CMC delineates discontinuous behavior in isotherms for the surface tension g, specific conductivity k, and turbidity t. For all three properties, the concentration dependence changes when the detergent concentration surpasses the critcal micellar concentration.
The surface tension decreases with increasing concentration of the surfactant indicating surface excess and monolayer formation. Above the CMC, however, the surface tension no longer changes, because the free monomeric concentration of the detergents remains constant. At any given concentration, detergent molecules are in euqilibrium between monolayer, monomeric solute, and micellar component. Only the number of micelles increases with increasing concentration of detergent above the criticall micellar concentration. This is evident by the increase in turbidity (light scattering) of the solution. If the dergent carries a [positive or negative] charge, the conductivity of solution increases with increasing free detergent molecules.
The observed discontiunity at the critical micellar concentration is itself a function of the size of the micelles formed. It is imporant to keep in mind that micelles have a defined size and aggregate number (number of monomers in micelle) and don't increase with increasing concentration, if the concentration does not dramatically exceed the CMC. The size and aggregate number of micelles, however, can be well characterized for different chemical structures of detergents.
The concentration above which micelles start to form is a function of monomer structure and chemical composition. The CMC decreases with increasing chain length of the apolar residue. The CMC decreases by about a factor of 0.3 when the hydrocarbon chain length is increased by two carbon atoms. Micelle size also increase with an overall increase in ionic strength of the solution. The observed conductance of ionic surfactants above the CMC is lower than for the monomeric species. This is mainly due to the presence of counter ions which interact more strongly with the more structured multiple charged micelle surface effectively shielding this charge. Micelles are only about 20% charged as compared to the monomeric species. This must clearly be an effect of supra molecular structure formation and charge-charge interaction between head groups as well as counter ions.
Table List of some commonly used detergents
for membrane protein studies
| Common name | Chemical name | mol. weight
(g/mol) |
CMC
(mM) |
CMC
(weight %) |
aggregation number per micelle |
| CHAPS
(zwitterionic, mild) |
3-cholamido propyl dimethyl ammonio-1- propane sulfate | 615 | 8 | 0.49 | 10 |
| SDS
(ionic, strong) |
C12-sulfate-Na+ | 288 | 8.2 | 0.24 | 62 |
| b-OG | C8-b-D- glucopyranoside | 292 | 25 | 0.73 | 27 |
| octyl-POE | polydisperse octyl oligo oxyethylene | 400 | 6.6 | 0.25 | 57 |
| TX-100 | tert. C8 phenyl poly ethylene glycole (9-10) | 628 | 0.24-0.34 | 0.015-0.02 | 100-155 |
Fig. Structures of CHAPS (top) and Triton detergents
from: A guide to the properties and uses of detergents
in biology and biochemistry; Calbiochem;
click here for PDF version of online
brochure;
Chaps and Triton X-100 are two very commonly used mild detergents for membrane protein purification. Unlike SDS ( a sulfonated alkyl unit) they do not denature most proteins and even leave quaternary or larger aggregate forms intact.
Light scattering
The size of ionic micelles as characterized by the number of surfactant molecules per micelle can be measured by light scattering experiments (van Holde, chapter 7.1), which allows a determination of molecular weight of particles in solution without specific information about their structure. Since visible light is electromagnetic radiation, the electric component of the wave should produce oscillation of the electron distribution of the targeted sample molecule. Electron oscillation induces dipoles which absorbe light energy while dispersing this energy in directions other then the direction of the incoming radiation. The oscillating dipole moment is a function of the molecular polarizability a of the molecule with the dipole moment m being the product of the polarizability a and the field strength E of the radiation.
m = aE
In general, we are inerested in the change in intensity that occurs in the direction of the incoming radiation I/i. This loss in intensity is proportional to the degree of scattering which is a function of the number and size of the particles (here micelles) in solution. Light scattering from a number of small particles (3-10 nm) as compared to the wavelength of the radiation (400-800 nm) is known as Rayleigh scattering. For all practical purposes, the polarizability of a molecule is a microscopic property and not easily determined for biological macromolecules. Rayleigh scattering theories related change in radiation intensity with the refractive index of a solution, a macroscopic property which is proportional to the polarizability as shown below. A convenient measure of polarizability for visible light scattering is the square of the refractive index n:
n2 - n02 = 4pNa
n is the refractive index of the solution, n0 the refractiv index of the solvent, and N the number of particles with polarizability a. It can be shown that scattering produced by a solution containing a weight concentration C of particle of molecular weight M depends on the product CM and light scattering measurements (Rayleigh scattering) not only indicate trubidity but can be used for molecular weight determination.
Distribution of micelles sizes is relatively homogeneous for any given detergent and the number of average aggregation numbers exhibits a narrow distribution. The current model of the structure of ionic micelles of synthetic surfactants in dilute aqueous solution is that of a rough-surfaced sphere or ellipsoid containing 40-150 monomers. The detergent most commonly used for the isolation of membrane proteins for analytical purposes is sodium dodecyl sulfate (SDS). It is a negatively charged, sulfar containing surfactant with a C12 carbon residue with a critical micellar concentration of 8.1mM.
Phospholipids are weak surfactants and have a very low CMC value because of their two long fatty acid chains. Due to their structure of a small head group with a large, bulky hydrocarbon part, phospholipids would ideally form elongated elliposidal structures. In fact, the ideal aggregate structure of phospholipids above their CMC are extended two dimensional sheet structures -- phospholipid bilayers. Phospholipids, however, solubilize very well in dergent micelles made from SDS.
The CMC for phospholipids has been measured by filtration techniques,
assuming that the phospholipid conentration of a solution pressed through
a paper filter absorbes more lipids in the aggregated form than the free
monomers in solution. Indeed, a discontinuity can be observed when comparing
the phospholipid concentration of the start solution with that of the filtered
solution above a certain concentration.
Below the CMC, original conentration and filtrate concentration are
equal, while above the CMC, much of the phospholipids are retained in the
filter because the 'micelle' aggregates get stuck in the pourous material.
The CMC for DPPC (dipalmitoylphsophatidylcholine) has been determined in
pure water as 0.5nM. The CMC increased when increasing the molar ratio
of methanol of the solution and reached 10mM in pure methanol indicating
good solubility of phospholipids in this organic solvent. Naturally occuring
phospholipids are thought to have CMC values in the range of 0.01 to 10nM.
This is six orders of magnitud lower than the CMC values for common detergents
used for membrane [protein] solubilization (for a further discussion on
phospholipids see section on vesicle preparation).
Isolation and purification of membrane proteins
There are many different detergents available for the solubilization of membrane proteins. Different classes of detergents are useful for different purposes and different membrane proteins. The most important factor in choosing a detergent is its effect on the protein structure. SDS is a strong anionic detergent which is primarily used for analytical procedures because it not only solubilizes membranes by forcing proteins and lipids into its micellar aggregates, but it usually denatures proteins by interfering with the hydrophobic packing of the core amino acid residues. In fact, SDS is the detergent of choice for analytical polyacrylamide gel electrophoresis (SDS-PAGE). For a protein isolated for functional studies, milder detergents have to be selected. A mild detergent is detergent that by definition does not cause a protein to denature or cause loss of function in vitro. The classes of detergents that compirse mild detergents are often non-charged and tend to have small aggregation numbers. In addition, detergent with small headgroup and long alkyl chains, and those with large head groups and short alkyl chains tend to be milder as shown here. See also table above.
Fig. Correlation between head group
and alkyl
side chain volume and 'mildness' of a detergent
Note: mildness refers to a the solubility property
that leaves the native protein structure intact; the
detergent
most suitable for protein solubilization has to be
determined experimentally and differs from protein
to protein; (from N.Koenig, University of Basel, Switzerland)
Some membrane proteins are so sensitive to detergent interaction that any loss of phospholipid-protein interaction causes a loss of function. The nicotinc acetylcholine receptor is one such protein. Other proteins are resistant against detergent denaturation, even in the presence of SDS. The outer membrane pore forming protein porin is a stable protein. It is a homotrimeric protein which only dissociates into its individual monomers in SDS at temperatures above 70 degrees Celsius. The SDS polyacrylamide gel below shows the result of a purification procedure of the general diffusion porin OmpF from Escherichia coli. The gel includes from left to right different fractions and samples of extracted E.coli membranes with increasing accumulation and puritiy of the OmpF porin. The native membranes (lane 1) contain a complex mixture of proteins, visualized by the many bands separated over the entire molecular weight range of the gel. With increasing purity and removal of undesired proteins by gel filtration and ion exchange chromatography, only a single protein species is left.
Fig. Purification of E.coli OmpF porin
std molecular weight standard, numbers indicate kDalton;
trimer -LPS porin trimer with bound lipopolysaccharide (LPS), a glycolipid
of bacterial outer membranes; monomer porin monomer; OmpA outer membrane
protein A;SDS-PAGE separates proteins according to molecular weight/charg
ratio. Small proteins travel far across the gel (top to bottom) while large
protein [complexes] mover slower and are retained. The molecular weight
markers are indicated on the far left of the gel ranging from 95kD to 36kD.
Purifed porin trimers (lanes A, B, and C run at an apparent molecular weight
of 95kD (trimer-LPS band), while the monomers, obtained by boiling the
porin-SDS extract, run at 35 kD.
Coomassie blue staining of proteins in polyacrylamide gels reveals the
banding pattern of proteins. This is due to differences in molecular weight,
quaternary structure (complex formation), or post translational modification. 
For
most proteins, subunit interaction is not strong enough to withstand the
denaturing effect of SDS. Bacterial porins, however, show an unusual resistance
toward SDS denaturation, including a strong association with the bacterial
cell wall glycolipid lipopolysaccharide (LPS). The small gel shown here
shows porin trimers in association with different amounts of the glycolipid
LPS. Control lane 1 shows LPS-free porin trimers, while the sample in lanes
2 and 5 contains porin trimers associated with native LPS from E.coli outer
membranes. Lane 3 shows a porin trimer with a synthetic, short chain LPS
exhibiting a low affinity. This lipid A type of lipopolysaccharide dissociates
from porin in SDS solution. Lane 4 shows a heat denaturated monomer. The
banding pattern in the right lane (m) contains the molecular marker proteins
as described above. The lower mobility of some porin trimers (banding pattern
with higher molecular weight) is due to the binding of different amount
of lipopolysaccharide units. The ratio of glycolipid to trimer is not well
defined and varies from sample to sample. Heat denaturation, however, removes
any residual glycolipids from porin monomers. Monomers never show a banding
pattern as do the native trimers. Below is a schematic representation of
the cell wall composition of E.coli showing major inner and outer membrane
components.
Abb.: LPS lipopolysaccharide; PL phospholipid; LPP Brown's
lipoprotein; X generic integral membrane protein; EnvZ regulatory protein
of the ompF - ompC operon; F0F1-ATPase ATP synthase of electron transport
chain; IM inner membrane; OM outer membrane; PG peptidogylcan; OmpA outer
membrane protein A;
Principles of gel electrophoresis
The principle behing the separation of proteins of different molecular
weight by polyacrylamid gel electrophoresis (PAGE) is based on the movement
of charged particles in the force field of an electric field. This can
in generally be treated as a transport process. Molecules can be separated
on the basis of their
- mass
- shape
- charge
for both analytical (very small amounts) or preparative (isolation
of large quantities of purified material) techniques. The transport process
is base on the equilibrium of two opposing forces that influence the particle
-- friction and force field (gravitation, voltage). The equilibrium of
the two opposing forces results in a constant velocity which is proportional
to one or a combination of the parameters mass, shape, and charge. A general
theory developed to understand the transport of molecules in aqueous solution
is called hydrodynamics. Stoke's law describes the relation between
the friction and viscosity h of
the medium:
fo = 6phR
with fo being the frictional coefficient and R the [Stoke's] radius of the particle. There exists no good theory for polyacrylamide gel electrophoresis, because the moving particle is immersed in a polymer matrix soaked in an electrolyte with SDS, in other words, a non-ideal solution. For the movement of a charged particle in an electric field, Coulomb's law is a good approximation for the force working on the protein:
F = zeE
with z the number of charges, e the unit negative charge of a single electron, and E the electric field strength in the gel. Since the electric and frictional force are equal at equilibrium (constant velocity of moving particle), the following relationship holds:
fn = zeE
The electrophoretic mobility U is defined as n/E = ze/f. For spherical particles the frictional coefficient can be replaced with Stoke's law:
z e
U = ------
6phR
Proteins are not separated due to their native charges alone, but strongly
associate with SDS molecules in an equal weight to weight ratio. Proteins
in SDS electrophoresis gels therefore carry negative charges proportional
to their size or molecular weight. The mobility is linear proportional
with the logarithm of the molecular weight. See the molecular weight standards
in the above polyacrylamid gels.
2D Gel electrophoresis
The charges of the amino acid residues are of course not umimportant and the net charge of proteins depends on the pH of solution. This pH dependency is used to determine the isolectric point of proteins and for protein separation techniques is known as isolectric focussing. Here, the proteins are separated in an electric fiel along a pH gradient. The proteins change their net charge and exhibit different charge/weight ratios as they move along the pH gradient. Thus a second dimension can be applied to separate proteins; the first dimension is the SDS associated charge/molecular weight separation; the second dimension the isolectric focussing of proteins of equal size in a pH gradient.
Fig. Schematic diagram of protein separation on 2D gel electrophoresis
2D gel electrophoresis allows the separation of proteins of equal molecule
weight but different charges in a pH gradient. Although developed in the
early 1970s, 2D gel electrophoresis was not widely used because of the
complexity of the banding (or spot) pattern on these gels and the limited
ability to unabmiguously identify which proteins belongs to which spot
on a gel. Technical information
on isoelectric focussing (pH gradient dimension) and molecular weight dimension
can be found at Expasy.
Proteomics
The dramatic advancement of molecular biology and the high thoughput
sequencing of complete genomes of more than 20 microorganisms and two eukaryotic
organisms (yest and C.elegans), 2D gel electrophoresis has experienced
a dramatic resurgence. 
The
new science of proteomics addresses the protein expression pattern of cells
to understand the response of cells or organisms for certain signals
-- hormonal, neurobiological, immunological, or developmental. Although
mRNA levels are normally used to assess the presence of proteins by studying
the gene expression levels (Northern blot analysis), many times the concentration
of mRNA poorly correlates with actual levels of proteins in a cell. The
image on the left is taken from a 2D gel of a plant cell extract from Arabidopsis
thaliana. As the sample gel shows, and this is only 10% of the entire pH
and molecular weight range on the original
gel (from 2D page museum at Expasy), there are literally hundreds of
spots from whole cell extracts. For each spot the proper protein or a post
translationally modified isoform (e.g. phosphorylation shifts charge/pH
dependency) has to be determined. This can be done by cutting out the piece
of interest, purifying the protein from the gel matrix and perform a biochemical
analysis like microsequencing, or simply checking for the presence of phosphorylation
or glycosylation. More sample pictures and detailed information about 2D
gel electrophoresis methods, analysis, and databases can be found at the
Swiss
2D Page server at Expasy.
Mass spectrometry
For whole cell analysis of 2D gels, determining the sequence of protein fragments extracted from individual spots by mass spectrometry is the method of choice. Mass spectrometry determines the charge/mass ratio which is higly significant for molecular structures. The smaller the molecule, the easier the interpretation of mass spectra. This is the main reason why for proteomics research, proteins extracted from 2D gels are fragmented into small peptides of 5-15 amino acids and the sequence including that of modified residues, is quickly determined by correlation algorithms that match charge/mass ratio with potential amino acid combination expected for the observed values. Phosphorylation adds two negative charges and an average of 79.9799 g/mol to the molecular weight of a peptide/protein. Partial sequences can then be pieced together and the longer (partial or complete) protein sequence used to search for homologuous proteins (or genes) using programs such as BLAST at GenBank of the National Center for Biotechnology Information, NCBI.
Current techniques of mass spectrometry (van Holde, chapter 5.4) used
for the elucidation of peptide sequences include MALDI (matrix assisted
laser desorption ionization) and ESI (electrospray ionization). In both
cases, peptides are ionized, accelerated in vacuum and the time needed
for them to travel over a specific distance correlated with their charge/mass
ratio.
 |
 |
The accuracy of mass spectrometry is better than for any other technique
(deviation from theoretical molecular weight less than 0.1%). Even though
the mass of large proteins can be assessed accurately, fragmentation of
proteins into small peptides is necessary when using the technique as a
micro sequencing device as mentioned earlier. The possible amino acid combinations,
let alone their sequence, can only be determined for short sequences by
matching the mass/charge ratio with possible amino acid combinations.
The PeptIdent server at Expasy contains a table
with experimentally determined mass values for all possible amino acids
and common chemical modifications. It is used to identify proteins with
peptide mass fingerprinting data, pI and Mw. Experimentally measured, user
specified peptide masses are compared with the theoretical peptides calculated
for all proteins in SWISS-PROT, making extensive use of database annotations.
6. Structure determination of membrane proteins
Circular dichroism to study secondary structures of proteins
Amino acids are optically active molecules and in a polypeptide are often found as part of regular secondary structures. The frequent occurance of alpha helices and beta sheets in proteins has thus been exploited by measuring the presence of such regularly arranged units from circular dichroism spectra of protein solutions. Although CD measurements are not useful to obtain high resolution structures, but merely secondary structure content of a protein, this information is useful to study the status of protein folds. It can generally be assumed that the absence of any alpha helical or beta strand components indicate the unfolded state of a protein.
Dichroism occurs when light absorption differs for different direction of polarized light. Light can be polarized either in a linear way, where the plane of the electric vector is fixed while its amplitude oscillates, or in a circular way, where the plane of polarization of the electric vector is modulated while the amplitude remains constant. The electric vector of circularly polarized light describes a helix which may be right-handed or left-handed.
Linearly polarized light will be absorbed maximally when parallel to the transition dipole of the sample molecule, while its amplitude is not affected when the plane orients perpendicular to the transition dipole. Transition refers to the transition between energy levels in a molecular structure of electrons in inner and outer shell, or their vibrational and rotational transitions. A transition has a measurable characteristic of direction, such as the different directions assigned to molecular orbitals, in which we can find electrons. This directionality associated with energy transitions can be exploited spectroscopically for structural information. This direction is called the transition dipole, not to be confused with molecular dipoles or induced dipoles for molecular interactions (see chapter 8.5 and 9.1.5 for details). The energy of a wave particel (photon) of different wave length interacts with different levels of interaction; as a consequence, microwave radiation is used to measure rotational transitions, infrared wave length yield information on vibrational transitions in molecules, while visible light und UV photon energies interact with electrons in the outer shell. This is the range of circular dichroism spectra used to determine secondary structure content of macromolecules. X-ray radiation is absorbed by electronic transitions of the inner atomic shell.
A molecule can be excited from a lower to a hihger energy state by the absorption of electromagnetic radiation. Microwaves thus increase the temperature of matter by increasing the rotational energy of molecules. Liquids are specially sensitive to energy absorption in the microwave range because the degree of freedom of individual solvent molecules. The increased rotational energy of water molecules, for example, contributes to their kinetic energy and thus increases the temperature of a cup of coffee.
In proteins the aromatic bond structures are of importance in spectroscopy. For protein structure studies we are primarily concerned with backbone conformations. The peptide bond amide group is the dominant chromophore of the polypeptide backbone and has a weak absorption maxima at 220nm and a stronger absorption maxima at 195nm. Circular dichroism makes use of the fact that right and left handed polarized light are absorbed slighly differently in asymmetric molecules. Even though individual amide groups in protein backbones have a symmetric transition dipole, their mutual interaction in highly oriented secondary structures induces asymmetries which translated into circular dichroism spectra (difference of absorption of left and right handed polarized light not zero). Protein secondary structure can be revealed based on their characteristic electronic circular dichroism behavior between 190 and 220nm (see Fig. 10.15 in van Holde for details).
Table Electronic circular dichroism pattern for protein secondary structures
| secondary structure | positive band at | negative band ata |
| alpha helix | 190nm | 222nm and 208nm |
| beta strand | 198nm | 215nm |
| beta turn | 210nm | 190nm |
An example of a peptide with a poly-(Valine-Lysine) sequence shows stabilization in beta strand conformation at pH2.3 after being incubated for several hours in 100mM NaCl solution (_____). The initial conformation indicates random coil structure (----). The prominent negative ellipticity at 215nm and a positive band at 195nm is typical for beta sheet formation.
Fig. poly (Val-Lys)n electronic CD spectra
This figure is taken from a study on peptide with alternating hydrophobic-hydrophilic amino acid residues and their possible significance for prebiotic evolution (A.Brack and L.E. Orgel, 1975, Nature 256,383). A random coil to beta sheet induction could also be achieved by adjusting the pH to 8.8. This transition was reversible and occured within 15 minutes. The peptide precipitated at pH 9.0 indicating the strong propensity of beta strand peptides to form elongated sheet structures. These sheets eventually become insoluble. Such a aggregation mechanism based on beta strand interaction is also suspected to be the major mechanism behind the formation of amyloidogenic plaques such as found in Alzheimer's disease.
Similarly, sedondary structure content of larger proteins can easily be determined using CD. This is a particularly useful technique to study folding behavior of proteins such as bacterial porins. These outer membrane proteins are composed of three identical subunit, each of which forms a 16 or 18 stranded anti-parallel beta barrel. CD spectra of porin preparations show typical minima at 215nm.
Fig. CD spectra of phosphate selective porin of E.coli
(PhoE; PDB accession number 1PHO)
and PDB structure of related OmpF at 2.4 A resolution (PDB accession number
2OMF)
curves from 1 - 5 represent decreasing urea concentration
(8M, 4M, 2M, 1M, 0.5M urea respectively). color code of secondary structure:
beta strand brown; alpha helix purple; random coil grey;
As shown in the figure for phosphoporin of E.coli (PhoE), CD
measurements can be used for protein denaturation and refolding studies.
Because porins are almost entirly made of beta sheet with less than 3%
alpha helical content, their electronic CD spectra are easy to interprete.
The figure shows a increase in negative ellipticity at 215nm with decreasing
urea concentration. Note that at 8M urea, porin proteins are completely
denatured.
Studying protein complex assembly - gap junction proteins
Porins are found in large copy numbers in the outer membrane of Gram-negative bacteria. A single E.coli cell contains as many as 105 units. From E.coli strains containing more than one porin gene like ompF, phoE, lamB which code for the general diffusion porin, the anion specific phosphoporin, or the maltose specific maltoporin, it can be shown that the porin content of the outer membrane can be changed in 10 to 20 minutes, demonstrating that these seemingly stable structures are highly dynamic. This is even more remarkable considering the fact that porin trimers are found in close proximity in the outer membranes, essentially forming a densly packes two dimensional moleculer sieve. This tendency of porin to form large aggregates or ensembles in the plane of the membrane, made its isolation and purification relatively straight forward, and also allowed the determination of its structure by electrons microscopy. Removal of phospholipids (and lipopolysaccharide) from outer membrane fragemnts results in almost pure, two dimensional porin crystalls.
Bacterial membrane proteins are not unique in forming this native two
dimensional sheets. Also muscular nicotinic acetylcholine receptor and
gap junction channels are found in 
large
plaques in specialized cell membrane areas of eukaryotic cells.
Gap junction channels are voltage gated, weakly selective pores with an
exclusion size limit of about 800 to 1200 Dalton. They form cell to cell
channels coupling the cytoplasmic compartments of neighbouring cells as
well as the membrane potentials of the bridged membranes. Schematic
diagram of gap junction channel: a connexon consists of six connexin
subunits and spans a single cell membrane. Two connexons of adjacent membranes
tightly bind together to form a 12mer connexin complex. Phospholipids (on
the left) indicate the membrane spanning domain (42Å) for each connexon
(70Å) bridging an extracellular gap between the two coupled membranes
of about 35Å. Inside the channel, two putatively charged rings (close
to extracellular side of each membrane) affect ion flux and selectivity.
The distance between these charged rings of residues is estimated to be
about 40Å. The small cylinder (dotted lines) in the upper connexon
indicates the location of an M3 transmembrane alpha helical segment (one
found in each subunit). Each subunit is predicted to have four transmembrane
spanning helices (M1 through M4), with M3 being an amphipathic alpha helix.
This alpha helical transmembrane domain structure has been confirmed by
the recently published electron diffraction structure of heart muscle connexin
Cx43 (also called a1). The C-terminal
end of each connexin is located at the cytoplasmic side and together with
a central cytoplasmic loop between M2 and M3 contributes 17 positive and
8 negative charges to the subunit surface.
Although no high resolution structure has been solved yet, a 7.5 angstron resolution from electron diffraction studies has been obtained and largely confirms the topology diagram discussed above.
Fig. 7.5 Angstrom map of a heart muscle gap junction (connexin
43 complex)
from Unger et al., 1999, Science 283,1176; see text for
details
The structure from electron diffraction studies of recombinant connexin 43 complexes shows the electron densitiy profile of an entire channel outlining the mostly alpha helical organization of the transmembrane spanning domains (M) and the molecular interface across the inter cell gap (E). The full view has been reduced to show the channle interior (panel b) and cross section of the image reconstruction map are shown in panel C.
This large protein complex spanning two cell membranes is particularly interesting in studying not only the structure function relation ship of the native channels, but also their synthesis, membrane assembly of a connexon in each cell and connexon-connexon assembly across the narrow gap of about 2nm between coupled cells as well as the large 2D sheet structures (plaques) that are formed in specialized membrane regions. .
Fig. Proposed assembly pathway for gap junctions
From Yeager, Unger, and Falk, 1998, Current Opin. Structural
Biology 8:517;
The figure above outlines the synthesis, assembly and intracellular transport pathway for gap junction channels in a hypotheticla mammaliona cell. Eight different stages of the process can be separated and studied independently -- synthesis, oligomerization, trafficking, intracellular storage, plasma membrane insertion, plaque formation, and two steps of degradation -- annular gap junctions and their [proteolytic] degradation in lysosomes and proteasomes.
How can such a pathway be elucidated? Here we concentrate on the formation of connexon structures, the hexameric 'hemi-channels' spanning single membranes. To determine connexon structures the following questions have to be answered: protein sequence and molecular weight of subunits, orientation of subunits in membranes, molecular weight and subunit composition of oligomers, and pathway and kinetics of trafficking to the cell surface.
The first step of connexon assembly in endoplasmatic reticulum membranes
(ER membranes) can be studied in an in vitro protein translation system.
These are cell free cytoplasmic extracts of red blood cells enriched with
membrane fractions (microsomes) of ER from dog pancreas. The latter is
specially rich in ER membranes because the main purpose of the pancreatic
cells is the production and secretion of hormones like insulin and proteases
like pepsin.
From Yeager, Unger, and Falk, 1998, Current Opin. Structural
Biology 8:517;
Using sucrose gradient centrifugation, protein particles can be separated according to their size in sucrose solution (see Zonal Sedimentation, chapter 5.2.2 vanHolde). The molecular weight of particles in these centrifugation experiments can be determined by determining the sedimentation coefficient S. Large coefficient indicate large complexes, low S values indicated small complexes. Since density is a measure of mass/volume, a tightly packed protein complex shows a higher density then a single monomer. This is shown in the gradient elution profile above. Radiactive labelled connexin a1 or b1 (and deletion mutants thereof DN) complexes are extracted from their cellular environemnent. Since alpha 1 (43kD) and beta 1 (31kD) connexins have a different molecular weight, they can be sepatated by SDS gel electrophoresis. The western blot analysis (using anti bodies specific against either connexin type) shows that both fraction from the centrifugation contain a mixture of the two connexin types. The electrophoresis analysis, however, does not tell us if they are found as complex or independent units.
Sedimentation is analogous to electrophoresis where molecules are moved
within solution at constant speed, pulled by a gravitational force and
opposed by a frictional force of equal but opposite sign. When the gravitational
force is generated in a centriguge, there are three forces acting on proteins
in solution:
- a centrifugal force Fc = w2rm
(due to angular velocity w, mass m, and distance r from center of rotation)- a bouyant force Fb = -w2rm0
(due to the density difference of the particel with mass m and the displaced solution with mass m0)- a frictional force Fd = -fn
With a constant angular velocity, the sum of all three forces will
be zero and the particle moving with a constant velocity n.
Fc + Fb + Fd = 0
For the mass of solution displaced m, we can substitute the product
of particle mass m, its partial specific volume n(-),
and solution density r. The sedimentation coefficient
S is defined by the velocity of the particel divided by the centrifugal
field strength (equation 1). A small volume of a protein solution put on
top of a sucrose gradient in a centrifuge tube will behave as a moving
boundary along the centrifuge tube axis, with the boundary broadening over
time because of diffusion of the proteins within solution. 
The
definition of the diffusion coefficient is given in equation 2 indicating
that diffusion is proportional to the kinetic energy of the particle and
inversely proportional to the frictional coefficient. Zonal sucrose
gradient centrifugation minimizes the inpact of diffusion making the moving
boundary more stable. If the protein solution contains a mixture of protein
complexes of different size, multiple moving boundaries will form after
several hours of running the centrifuge. In addition, the boundary zone
for large particle will be sharper because diffusion is much smaller due
to the larger frictional forces of the large particles as compared to smaller
ones. Although the sedimentation coefficinet is a characteristic (intrinsic)
property of each protein, nucleic acid, polysaccharide etc., it depends
on the temperature and ionic strength of the solution. Experimental measurements
of sedimentation coefficient S therefore are related to standard conditions,
for which a characteristic S20,w
has been determined (tabulated in many references). Finally, S will also
depend on the particel concentration because particles may interact with
each other resulting in an apparent increased frictional coefficient.
When f0 is the frictional coefficient at concentration zero
(ideal solution, no particle interaction), the effective sedimentation
coefficient S is determined as indicated by equation 3 (with S0
the sedimentation coefficient for an ideal, diluted solution at C = 0).
Sedimentation coefficients are larger for large particles (see 9S and 5S particle for connexin hexamer and monomer, repsectively). Because the sedimentation behavior is not only a function of size, but also shape, the relation ship between sedimentation coefficient and molecular weight is not staightforward. The Svedberg equation (equation 4) shows how the molecular weight M of a protein can be determined by measuring both S and D of the protein in solution. Idealy, if a protein is globular and not hydrated, a plot of M versus S20,w yields a linear line. In fact, plotting S values of proteins this way is a fast way of determining if they are globular or not.
In a sucrose density gradient, the velocity of the moving particle will
continually decrease when moving into an increased sucrose concentration.
This, in fact, allows the separation of macromolecules and complex particles
like membrane of different origin with different lipid to weight ratio
not according to the size, but their apparent density. If the sucrose gradient
at its highest density is higher than the density of any particles in solution,
the velocity will be so slow that the centrifugation separates particles
of different density at equilibrium conditions.
GFP - green fluorescence protein
Fluorescence labelled antibodies (FITC
- Fluorescein Isothiocyanate - label on secondary antibody, binds
to primary antibody which specifically recognized a target protein)
have 
been used
for a long time to show the cellular localization of proteins. Also the
addition of smaller fluorescence labels has been employed that reversibly
bind to macromolecules or can be chemically cross linked to demonstrate
the surface exposure of certain amino acid residues. Recently, autofluorescent
proteins have been used through recombinant DNA technology to avoid the
often chemically harsh treatment of cells neede to bring fluorescence labels
into subcellular compartments. Here, protein chimeras are made where a
protein 'gains' a domain which carries a label which can easily observed
by light microscopy. One very successfull label protein is the green fluorescent
protein, or GFP.
A 236 amino acid, beta barrel forming protein from the jelly fish Aequorea Victoria has revolutionized 'live' imaging of cellular processes. Green fluorescence protein -- GFP -- exhibits fluorescence in the green visible range after being excited by blue light (fluorescence emission wavelength are longer that excitation wavelength). The three consecutive amino acid residues -Ser65-Tyr66-Gly67- inside the beta barrel structure autocatalytically combine to form a chromophore.
Fig. GFP protein: side and top view of barrel with central chromophore
at 2.13 Å (1EMB)
Fig.Composition of GFP chromophore (link)
and rasmol version (from PDB accession 1EMB)
The final fluorophore contains a series of "conjugated
double bonds" (ie. an alternating series of single and double bonds)
that results in the fluorescent properties of GFP. Chromophore name: DECARBOXY(PARAHYDROXYBENZYLIDENEIMIDAZOLIDINONE)
THREONINE
Fig. GFP is a 236 amino acid protein with the following sequence:
>1emb_ mol:protein-het length:236 Green
Fluorescent Protein (Gfp)
MSKGEELFTG VVPILVELDG DVNGHKFSVS
GEGEGDATYG KLTLKFICTT 50
GKLPVPWPTL VTTFXVQCFS
RYPDHMKRHD FFKSAMPEGY VQERTIFFKD 100
DGNYKTRAEV KFEGDTLVNR IELKGIDFKE
DGNILGHKLE YNYNSHNVYI 150
MADKQKNGIK VNFKIRHNIE DGSVQLADHY QQNTPIGDGP
VLLPDNHYLS 250
TQSALSKDPN EKRDHMVLLE FVTAAGITHG
MDELYK 236
The X in the sequence above indicates the chromophore position.
Fluorescence, general
Fluorescence occurs when excited electrons in a singlet state (electron
spins in molecule are 
paired;
see orientation of arrows in picture), after absorption of light quanta,
fall back into their ground state and emitt themselves light in the visible
range. The wavelength of the emitted light is usually longer than that
of the activating light because of internal loss of energy before fluorescence
occurs. This phenomenon is called the Stoke's shift and refers to the loss
of energy during the excited state of the electron (no radiation) where
internal conversion brings the excited electron to the lowest (first) level
singlet.
EX (to 2nd or higher singlet) excitation -
excitation state lifetime - emisson (from 1st singlet)
EM
While absorption is very fast and takes about 10-15 seconds, internal conversion to the first singlet level is about 1,000 fold slower. The actual lifetime of the electron in the first excited singlet, before fluorescence emission occurs, is fairly long and lasts about 10-8 seconds. Fluorescence is a spontaneous process and is favoured in conjugated double bond systems of the pp* transition, because the probability of a spontaneous emission is related to the integrated intensity of an absorption band; the stronger the intensity of absorption of the first singlet, the higher the probability for fluorescence (the energy of a transition is determined by the spacing between the energy levels; however, not all photons of the correct energy quanta will be absorbed, the fraction of photons absorbed is the intensity of absorption; see vanHolde chapters 8.4 and 11 for details). This is found in sidechains of tryptophane and tyrosine amino acids and is more pronounced in the autocatalytically derived chromophore sidechain of GFP.
The entire wavelength of visible light results in a fluorescence spectra,
either absorption or emission spectra. Excitation of a fluorophore
at three different wavelengths (EX 1, EX 2, EX3) does not change the emission
profile but does produce variations in fluorescence emission intensity
(EM 1, EM 2, EM 3) that correspond to the amplitude of the excitation spectrum.
Quenching
Since loss of energy of an excited electron can occur as radiation (fluorescence)
and non-radiation process (internal conversion, heat), the quantum yield
of a substance is the ratio of the total number of quanta emitted to the
total number of quanta absorbed. The quantum yield can be related to the
loss of fluorescence over time, or fluorescence decay, which follows
a first-order process, like radioactivity. In other words, the fluorescence
events of each molecule is independent of its neighbors. The number of
molecules N(t) that still can emitt fluorescence after time t is given
as:
N(t) = N(0) e-kt
By definition, the lifetime t of an ensemble of N fluorescent molecules is 1/k. The lifetime can be measured because the intensity is proportional to the number N of molecules in the excited state at time t. The lifetime is simply the time it takes for the maximum intensity to decrease by the factor 1/e.
The quantum yield differes with the molecular environment because this
determines the potential non-radiation loss of energy. The quantum yield
of a fluorophore often increases when the molecule is taken up from solution
and binds to DNA or proteins. Thus, measuring quantum yields is indicative
of binding. It is not necessary to know the exact quantum yield in order
to use fluorophores for biochemical studies. The intensity of the fluorescence
spectra changes with changing environment (solvent, ionic strength, lipid
environment, pH etc.) and is referred to as quenching if the intensity
decreases. The loss of intensity will depend on fluorophore concentration,
shape of fluorescence spectrum and the wavelength chosen for observation
(intensities vary differently at different wavelength). This is of course
reminiscent of electronic CD spectroscopy.
Energy transfer
A special form of quenching is the energy transfer between fluorescent molecules, where a donor activates electrons in an acceptor molecule. This interaction is a function of the distance between donor and acceptor and can be exploited to study conformational flexibility in macormolecues. The distance dependency follows an r-6 function and thus is a short range effect. Requirements for energy transfer are transition dipole interaction between the two fluorophores and an overlap of the fluorescence spectrum of the donor with the absorption spectrum of the acceptor.
Photobleaching
Under high-intensity illumination conditions, the irreversible destruction or photobleaching of the excited fluorophore becomes the factor limitingfluorescence detectability. Recent investigations have provided a detailed description of the multiple photochemical reaction pathways responsible for photobleaching of fluorescein (excitation/emission maxima ~494/520 nm; spectra). Some pathways include reactions between adjacent dye molecules, making the process considerably more complex in labeled biological specimens than in dilute solutions of free dye. In all cases, photobleaching originates from the triplet excited state, which is created from the singlet state via an excited-state process called intersystem crossing. (from Molecular Probes -- Introduction to fluorescent techniques). Fluorophores can be produced that show remarkable stability during long exposure to light (see movie on improved photostability of Alexa Fluor 488 phalloidin).
For links to see the applications of GFP in cell biology research,
single molecule detection and general information about fluorescence spectroscopy,
follow these paths:
|
GFP links |
Single Molecule Spectroscopy |
Fluorescence (general) |
| The laboratory of Kevin
Sullivan at The Scripps Research
Insitute
A microscopy page at the University of Pennsylvania Fluorescence images of connexin-GFP constructs (Matthias Falk, The Scripps Research Institute)
|
Single molecule spectroscopy
(ETH Zuerich, Switzerland)
Recording
(3.4M movie) of F1-ATPase single molecule motor (Montemagno
Lab, Cornell University)
|
Molecular Probes, a company the
provides almost every kind of fluorescence label, which provides an introduction
to fluorescence spectroscopy
Photobleaching
effect (Molecular Probes)
|
7. Functional membrane protein reconstitution
Membrane vesicles
To study the function of membrane proteins, they need to be in a membrane or phospholipid bilayer. Their are two types of artificial bilayer types -- spherical and planar. The first are referred to as vesicles or liposomes and the latter as planar lipid bilayer or black lipid membranes (BLM). Vesicles contain a lumenal aqueous compartment which is separated by the membrane. Membrane proteins incorporated into vesicle membranes, therefore, can be studied for transport processes.
Vesicles come in two forms -- unilamellar and multilamellar. Only the unilamellar vesicles are of interest here because they contain exactly one single membrane, where as the other types contain multiple layers and are not suitable for membrane transport studies.
Fig. Unilamellar und multilamellar membrane vesicles
Unilamellar vesicles belong to two categories, small vesicle called
SUV, and larger vesicle called LUV. Small unilamellar vesicle have a diameter
which is usually smaller than 200nm. It is these kind of SUVs that are
the preferred vesicle species used for transport studies, and as we will
see later, for the formation of planar lipid bilayers.
The formation of vesicles is achieved by a few widely used techniques:
- sonication
- freeze-thaw cycles
- detergent dialysis
While the first two techniques are fairly rapid (several minutes),
detergent dialysis requires several hours or days depending on the detergent
type, i.e., CMC value. Sonication makes use of high energy ultra-sound
water bath preparations. The sample, a phospholipid water solution is put
in the center of a sonication chamber. The lipid solution quickly turns
translucent indicating the formation of vesicles. The clearer the solution
(see light scattering), the smaller the particles in solution. It depends
on the protein used for reconstiution if sonication is an appropriate method.
Often, larger vesicle fragments are decreased in their size by brief pulses
of ultra sound sonication.
Freeze-thawing of lipid solutions makes use of the phase behavior of phospholipid layers. Freezing induces crystalline layers which can break into smaller fragments. These fragments start to form vesicular structure after repetitive freezing and thawing of the lipid solution. The advantage of both sonication and freeze-thawing is the absence of detergents at any time during the procedure. Native membrane fractions can thus be reduced in size on isolated as SUV preparations.
Detergent dialysis is the method of choice for complete control of vesicle
type (SUV) and composition. Here, membrane proteins and phospholipids are
solubilized in detergent extracts 
in
the form of mixed micelles. Placing the detergant extract solution into
a small volume dialysis bag, which in turn is placed into a large volume
of buffer (no detergent, no lipids, no proteins), the free monomeric detergent
will slowly diffuse into the large volume, while micelles are too big to
cross the porous material of the dialysis membrane. The phospholipids will
be retained inside the dialysis bag because their CMC value is severla
orders of magnitude lower than the CMC of the detergent. Hence the removal
rate of phospholipid monomers is so slow that it can be neglected over
the duration of the dialysis. When the detergent concentration inside the 
dialysis
bag reaches the CMC value or drops below it, the phospholipids, together
with the proteins, will assemble into SUV structures. Depending on the
protein, they are oriented asymmetrically (triangle indicates structural
asymmetry; arrow indicates functional asymmetry, i.e., active transport
in one direction such as for proton pumps bacteriorhodopsin or H-ATPase;
ion flux through ion channels has no intrinsic asymmetry) or are found
in a 50-50 or random orientation. Adjusting the lipid-protein ratio before
dialysis, the number of proteins per SUV can be estimated, allowing for
the control of membrane protein assembly.
Large unilamellar vesicle are used for electron microscopy studies.
If the lipid-protein ratio is very low (as low as 1:1 molar ratio), regular
2-D structure are obtained suitable for electron diffraction studies. The
regularity of these protein sheets can be enhanced by further removing
lipids using very carefully controlled detergent extrection of residual
phosopholipids (a process called delipidation).
Planar lipid membranes
Planar lipid bilayers have been developed for the sole purpose of studying ion channels in a very simple system and with as few components as possible. Planar bilayers allow a choice of lipids, lipid composition, membrane protein, protein orientation, and easy adjustment of electrolyte conditions on each side of the bilayer and total control over the transmembrane potential.
The basic setup for bilayer fomation as developed by Maurice Montal (UCSD) and enhanced to a solvent free system by Hansgeorg Schindler (University of Linz, Austria) is shown below:
Fig. Planar lipid bilayer formation from monolayer assembly
from H. Schindler, 1980, FEBS Letters 122:77-79
A teflon two-chamber module is filled with vesicle solution and separated above the water levels of these solutions by a thin teflon foil or septum (12mm) which contains a single aperture with a diameter between 50 - 300mm.
Fig. Teflon septum 'sandwich'
The inner circle (light yellow) represents the 12mm
teflon foil and contains a punctured aperture of 100mm
(not visible in this picture). The central thin foil (yellow) is stabilized
between two circular sheets of thick teflon foils (light gray) that each
contain a cutout circle of about 3-5mm exposing a small area of the thin
foil. Note that the thickness of a bilayer is 50nm and thus about
240 times thinner than the supporting thin foil.
Read more on how to make a cell membrane!
Monolayers will spontaneously assemble from vesicles in solution. Alternatively, a hexane/lipid droplet can be place onto each half cell solution. While hexane quickly evaporates, phospholipids will form stable monolayers that show an liquid phase L2 surface-pressure area behavior. A few minutes after placing the vesicle solution into the teflon chamber, monolayers are stable enough and will form a bilayer across the small aperture in the teflon foil separating the two half chambers when the two solutions are carefully rised above the septum aperture.
The monolayers formed from SUV solutions are in equilibrium with the vesicles. The surface pressure, called equilibrium pressure pe, of the monolayer thus formed is a function of the SUV diameter.
Fig. Correlation between SUV diameter and monolayer surface pressure
from H. Schindler, 1980, FEBS Letters 122:77-79
Proteins can be inserted into the membrane before or after bilayer formation. The figure on bilayer formation shows that proteins are already incorporated into the vesicle membranes. This means that membrane proteins that will be part of the newly formed bilayer structure will temporarily be inserted at the air-water (or teflon-water) interface and are susceptible to denaturation. The use of this technique, however, demonstrated that many membrane proteins reconstituted from vesicles into planar membranes are functional, although these studies are not quantitative and the observed activity is focussed on a very small number of active proteins only, usually in the range of one to less than one hundred. The goal of these membrane electrophysiology studies in most cases are so called single channel recordings, which depend on the presence of only one single channel protein complex.
Monolayer experiments can show the degree of protein deanturation of membrane proteins at the air water interface. As discussed earlier, p-A isotherms are obtained under equilibrium conditions and the path of the isotherm is thus independent of the direction of the change in surface area. Monolayer containing membrane proteins, such as native membrane vesicle preparations often show non-reversible isotherm behavior.
Fig. p-A isotherms from postsynaptic
vesicles (contain nicotinic acetylcholine receptors)
from Schuerholz and Schindler, 1991, Eur.Biophys.J. 20:71-78
The figure shows that repetitive compression relaxation of surface area results in increasing minimal area/molecule values. The first isotherm obtained shows a minimal A value of about 150Å2, the second of about 200Å2, and the third cycle shows a minimial A value of about 300Å2. This increase can clearly be attributed to the denaturation of proteins at the interface. It can be shown, however, that keeping a minimal surface pressure above 15mN/m inhibits the denaturation of most membrane proteins, including the nicotinic acetylcholine receptor shown above (simply stay within the cyclic isotherm on the very left). Since the equilibrium pressure of the vesicles used for bilayer formation is around 30mN/m, this reconstitution technique provides a safe and solvent free way of incorporating membrane proteins into planer lipid membrane systems.
Alternatively, planar membranes can be formed from pure lipid vesicles only and membrane proteins are incorporated after bilayer formation by either vesicle fusion or addition of detergent extracts to one side of the bilayer. The latter is a suitable method when using mild, non ionic detergents like Triton X-100 or beta-octyl glucoside at concentrations right above their CMC value. This keeps the membrane proteins soluble, and after adding a few microliters of detergent/protein extracts to the 1ml volume of these membrane system, the detergent is instantly diluted far below its CMC value and spontaneous incorporation of membrane proteins can be observed. These detergents at concentrations below the CMC do not destabolize plananr bilayers nor do they induce channel like pores that intefere with ion channel measurements.
The formation of these bilayers can not be monitored visually because they are too small. The first bilayers, however, have been formed by monitoring light refraction of a decane/lipid droplet covering the septum aperture ('painted' membranes). The slow diffusion of the decane into bulk solution resulted in a thinning phospholipid layer eventually forming a bilayer. The decreasing solvent content of the droplet diminished light reflection and the black picture was an indication of completion of membrane formation, thus the name black lipid membranes or BLMs. This technique is still in use because it is dramatically less challenging than the vesicle supported bilayer technique developed by H. Schindler. The substantial residual hydrocarbon in the membrane, however, may affect channel activity. The whole process of solvent free membrane formation is followed by measuring decrease in impedance or increase in AC current once the two half chambers are in contact through their respective monolayers (note that when the bilayer breaks or does not form, the current signal is in saturation. i.e., short circuiting the electrodes). The successful formation of a bilayer can be seen below (left panel) when observing the increase in A/C current across the teflon septum to a maximal level indicating that the entire aperture is covered and the size of the membrane does no longer increase.
Fig. Bilayer formation followed electronically (AC current) and
ion channel recordings (DC current)
from Schindler and Feher, 1976, Biophys.J. 16:1109-1113
When ion channels are incorporated into the membrane, rectangular jumps in DC currents are indicative of opening and closing transitions of ion channels (see part III for more details). The recording shown here were obtained from Gramicidin A peptide solutions.
