`sowmrnr as A runcnou or mum
`smuaun: Ann sown" COMPONENTS
`Catherine H. Schein
`
`?!$&&1!)+9;7.!*;,218014/!(75;6!!0996’##<<<"4+9;7."-53#4+9;7.,159.-04525/=
`
`Department of Organic Chemistry, CHN E56, Swiss Federal Institute of Technology, CH8092 Zurich, Switzerland.
`
`
`This review deals with ways of stabilizing
`proteins against aggregation and with
`methods to determine, predict, and in-
`crease solubility. Solvent additives (osmo-
`lytes) that stabilize proteins are listed with
`a description of their efi'ects on proteins
`and on the solvation properties of water.
`Special attention is given to areas where
`solubility limitations pose major prob-
`lems, as in the preparation of highly con-
`centrated solutions of recombinant pro-
`teins for structural determination with
`
`NMR and X-ray crystallography, refold-
`ing of inclusion body proteins, studies of
`membrane protein dynamics, and in the
`formulation of proteins for pharmaceuti-
`cal use. Structural factors relating to sol-
`ubility and possibilities for protein engi-
`neering are analyzed.
`
`t is generally known that proteins must be stored in
`an appropriate temperature and pH range to retain
`activity and prevent aggregation. This review dis-
`cusses varying other solvent properties to maintain
`the stability and solubility of proteins.
`Proteins are often most soluble in solution conditions
`
`mimicking their natural environment. Serum proteins are
`soluble in a pH and salt range where mature insulin,
`which is stored in acidic granules in the cell, precipitates‘.
`Bacterial proteins may prefer buffers containing gluta-
`mate or betaine, compounds that accumulate in response
`to high concentrations of Cl‘ in the medium? Caseins
`and other Ca2+ associated proteins may require small
`amounts of this ion to maintain their native structure
`
`during purification”. The stability of lactase (B-galacto-
`sidase) is greatly increased in the presence of milk pro-
`teinss. But for most proteins, experimental determination
`of the solution properties can help in solvent design.
`Low solubility in aqueous solvents is often regarded as
`an indication that a protein is “hydrophobic”, as aggrega-
`tion of integral membrane proteins after transfer to a
`hydrophilic environment is a well described phenome-
`non7. But all proteins are to some extent hydrophobic,
`with tightly packed cores that exclude water”. As native,
`properly folded structures aggregate less than unfolded,
`denatured ones, there is an intimate relationship between
`solubility and stability. The free energy of stabilization of
`proteins in aqueous solution is very low (ca. 12 kcal/mole
`at 30°C”); consequently proteins totter on the verge of
`denaturation‘oi“. Protein stability can be increased by
`solvent additives or by alteration of the protein structure
`itself.
`
`THE PROPERTIES OF PROTEINS IN SOLUTION
`
`Defining solubility. The chemists definition of solubili-
`ty, parts purified substance per 100 parts pure water, is
`not useful in a biological frame, as proteins in nature are
`never found in pure water. Blood and eukaryotic cyto-
`plasm contain on the order of 0.15 M salt, with large
`quantities of trace metals, lipids and other proteins. The
`cytoplasm of bacteria is more variable, with salt concentra-
`tion ranging from 0.3—0.6 M2. The solubilizing effects of
`small molecules and even other proteins means that
`protein solubility does not correlate with purity”.
`Operationally, solubility is the maximum amount of
`protein in the presence of specified co-solutes that is not
`sedimented by 30,000 g centrifugation for 30 min”. An
`even stricter criterion, function retained after centrifug—
`ing for l h at 105,000Xg, has been suggested for mem-
`brane proteins”. If one has a pure, lyophilized protein or
`a salt precipitate, one can determine solubility by adding
`increasing amounts of weighed solid, centrifuging, and
`measuring the protein content of the supernatant. Dis—
`solved protein should reach a maximum (maximum sol-
`ubility) and level off. (However,
`in the food industry,
`solubility is defined by sediment (in ml) remaining after
`centrifuging; the solubility index is thus inverse to the
`actual solubilitle.)
`The method described in the heading of Figure 1 allows
`definition of the solubility range of a protein in solution.
`A protein solution is diluted into a buffer series and the
`samples centrifuged in microconcentrators. As one can
`conveniently concentrate about 50 fold, a relatively small
`amount of protein is sufficient for the estimation.
`Measuring stability. Methods for determining the ther-
`modynamic stability of proteins use pH and temperature
`extremes or high concentrations of denaturants‘o. Al-
`though useful for discerning changes in the structural
`stability of mutant proteins that are not clear from activity
`data, they are not directly correlatable with the half life of
`proteins in solution. Since aggregation occurs at tempera-
`tures well below the Td for proteins, additives that stabi-
`lize proteins against aggregation may not necessarily af-
`fect the Td”.
`
`The major problem with using thermodynamic mea—
`surements is their failure to account for the kinetic effects
`
`that lead to aggregation. Both the enthalpy (AH) and
`entropy (AS) of hydration vary greatly with temperature,
`but they cancel to give a relatively small measured free
`energy (AG) of hydration that seems to vary little with
`temperature. Most of the temperature dependent kinetic
`contribution, which is the more important in explaining
`hydrophobic effects, dissipates in alterations of the solvent
`structure around the protein and reversible deformation
`of the protein structure itself”-15. Accurate determination
`of hydration shells can only be done from crystal struc-
`tures. Clearly other methods of determining protein sta-
`bility are needed.
`Proteins with shorter half lives generally have larger
`subunit molecular weight,
`lower isoelectric points (pl),
`higher affinity for hydrophobic surfaces, and greater
`
`
`
`
`
`
`
`BIO/TECHNOLOGY VOL8
`
`APRIL 1990
`
`
`
`AMGEN INC.
`
`Exhibit 1022
`
`Ex. 1022 - Page 1 of 10
`
`AMGEN INC.
`Exhibit 1022
`
`

`

`olishing Group http://www.nature.com/naturebiotechnology
`?!$&&1!)+9;7.!*;,218014/!(75;6!!0996’##<<<"4+9;7."-53#4+9;7.,159.-04525/=
`'rotein A
`+ Protein 8
`
`
`
`dues at the protein surface. Proteins are applied in high
`salt (0.7—1 M ammonium sulfate), which furthers hydro-
`phobic interactions, and then eluted with a decreasing salt
`gradient. Most proteins elute between 0.5 and 0.1 M salt;
`very hydrophobic proteins will not elute into low salt
`buffer unless the polarity is decreased by adding ethylene
`glycol. If a protein does not bind to Phenylsepharose, it
`either has a very hydrophilic surface (eg, RNase A) or it is
`aggregated.
`One can determine the hydrophobicity of a purified
`protein or follow changes in exposure of hydrophobic
`groups during folding by measuring interaction with a
`hydrophobic dye or radioactive tracer (eg,
`l-anilino—8~
`napthalenesulfonate‘“, or 125I-TID, 3-(triflu0romethyl)—3-
`(m-(125)iodophenyl)diazirine3).
`Aggregation and precipitation. Precipitation via any
`agent can be: (I) Reversible, as after precipitation with
`salts or large organic molecules like polyethylene glycol
`(PEG). Because PEG molecules are excluded from the
`surface of the protein, a two phase system develops and
`the protein is concentrated into a smaller volume, where
`its chances of interacting with another protein molecule to
`form an aggregate are increased (“excluded volume”
`modelzzv”). When the precipitant is removed, the water
`layer around the original molecule can reform and the
`protein molecules separate into soluble monomers.
`The protein structure does not significantly change
`during reversible aggregation. A plot of protein in solu—
`tion versus the concentration of the precipitant should
`look the same whether it is made with increasing precipi—
`tant‘(to precipitation) or decreasing precipitant (to solubil-
`ity). Reversibility is assumed for most mathematical mod-
`els of salting out” as well as some recent models of low salt
`aggregation phenomenai’mfi.
`(2) Partially reversible, a behaviour frequently seen in
`pH induced precipitation. Proteins precipitate around
`their p1 and resolubilize as the pH is adjusted upward or
`downward. But during the pH adjustment, residues may
`change orientation. When the pH is readjusted, they may
`not be able to regain their former position and a mixture
`of structures (isomers) results. Even primary structure can
`change if a protein is held at acid pH for long periods of
`time, as for example the deamidation of asparagine 21 of
`insulinl. A plot of protein in solution as a function of pH
`will depend on whether the protein has already precipitat-
`ed. Kinetic modeling of pH dependent aggregation has
`been attempted by linear regression”. Models could also
`use hysteresis equations.
`(3) Irreversible, which is usually initiated by extreme
`changes in the solvent leading to protein denaturation.
`But some proteins (Fig. 1) also precipitate irreversibly
`when concentrated above their maximum solubility in a
`given buffer. Inactive flakes of protein form and remain
`insoluble even on redilution of the sample or transfer to a
`buffer of the correct salt concentration. The nature of this
`
`tight intermolecular binding is not easy to study, as the
`aggregates arise from many-body interactions potentially
`involving all parts of the protein. The initiation could be
`direct interaction of surface hydrophobic residues, or, as
`aggregation shows cooperativity, partial disturbance of
`the hydration sheath or unfolding of the protein structure
`allowing interaction between normally “buried” residues.
`Irreversible protein aggregation is not easily modeled.
`Thermal denaturation curves are done at very low protein
`concentration, to avoid aggregation terms in the equa—
`tionsl".
`BUFFER DESIGN FOR MAXIMIZING SOLUBILITY
`
`
`
`
`
`Proteinconcentrationinsupernatant,mg/ml
`
`o
`
`200 400 600 8001000120014001600
`
`KCI concentration (mM)
`
`m I Solubility of T7 RNA polymerase (T7RP) as a
`function of salt concentration in 10 mM Na cacodylate buffer,
`pH 7.0,
`1 mM DTT, and 0.02 mM PMSF. The polymerase
`solution (ca.
`1 mg/ml in 0.1 M KCl, 20 mM tris pH 7.9, 5%
`glycerol) was diluted 1:10 with the indicated buffers and each
`sample was individually concentrated in 30 kD MW cutoff
`“Centr‘icons” (Amicon). The protein in the supernatant (mea—
`sured by the Coomassie blue assay) after concentration is
`indicated. The top curve (“Protein B")
`is from a second
`measurement using a finer salt gradient and more protein per
`sample.
`
`susceptibility to proteases. Both of the latter characteris-
`tics can be used as the basis for determining enzyme
`stability in less extreme environments as well as the effect
`of additives on stability. As less stable proteins have a
`higher tendency to adsorb to surfaces”, resistance to
`mechanical shaking may be a useful indicator of solution
`half-life‘s. Trypsin digestion has been used to define the
`salt stabilization of hyalin5.
`Determining surface charge. Isoelectric focusing gives
`the pl, the pH at which the protein shows no net charge in
`isoionic conditions. However, due to the binding of salt,
`one cannot assume that a protein in solution will be
`negatively charged at pH’s above its pl (eg, acidic caseins
`bind Ca2 and appear positively charged at pH 7“). At pH
`7.5 and 50 mM salt, most proteins will bind to DEAE—
`coupled resins if they are negatively charged and to
`phospho- and other negatively charged resins if they are
`positively charged. The charge strength can be estimated
`from the salt concentration required for elution. Gel
`methods for following the changes in surface charge
`during protein folding and aggregation have also been
`developed‘g.
`Generally, charged proteins can be “salted in” by
`counter ions. Binding of salts to proteins decreases bound
`water as well as the net charge at
`the surface. The
`solubility of lysozyme, a positively charged protein, was
`shown to vary more with the anion added than the cation;
`the anion dependence followed the Hofmeister series”.
`The solubility of caseins with pI between pH 3 and 5
`varies with the cation: sodium, potassium and ammonium
`caseinates are all more soluble than those prepared with
`calcium or aluminum“?
`
`Determining hydrophobicity. Binding to resins cou-
`pled with hydrophobic groups,
`like Phenylsepharose
`(Pharmacia) indicates the presence of hydrophobic resi-
`
`The properties of water as a solvent. Water's high
`dielectric constant and its tendency to solvate ions makes it
`an active copartner in enzyme reactions. When NaCl
`
`BlO/lECHNOLOGY VOL 8 APRIL 1990
`
`
`
`Ex. 1022 - Page 2 of 10
`
`

`

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`
`crystals are added to water, the atoms attract each other
`with only about 1/80th of the force in the dry state and the
`crystal dissolves. Analogously, dissolved proteins are coat—
`ed with a “hydration shell” around charged and polar
`groups that prevents self-binding. This bound water does
`not freeze (some proteins are even efficient antifreezes“) '
`and has different properties from the surrounding sol-
`vent molecules”. Bulk water molecules and the protein
`are in continual fluctuation, which leads to instability in
`the system11-30‘3‘.
`Protein stability in the solid state vs. solution. On the
`other hand, a protein completely stripped of its hydration
`shell is difficult to redissolve, as intermolecular hydropho-
`bic forces must be broken. Lyophilization and other
`drying methods should thus be used with caution and
`osmotic stabilizers added where necessary to insure that
`the protein can be rehydrated. The water content of dry
`milk powder is a compromise between shelf life, which
`decreases with increasing content of water, and solubility,
`which increases with hydration index”.
`Proteins in the solid state have different
`
`levels of
`
`reactivity depending on the water content. Dried protein
`with a water content below 22—25%, the minimum re-
`quired for conformational flexibility and activity, is ther-
`mostable. Glycerol, which stabilizes proteins in solution,
`acts as a humectant on the powder and causes decomposi-
`tion (as indicated by the Maillard browning reaction) at
`much lower water content (5—l5%). Conversely, sorbitol
`competes for the hydration water of the protein and does
`not enhance denaturation”.
`
`Solvent additives. There are many potential stabilizing
`co-solutes for proteins (Table l). Buffers are described in
`several excellent reviewsl’i’»33 and will not be covered here.
`
`Table 1 is separated into groups of compounds that have
`varying effects on the solvation properties of water: the
`dielectric constant, chemical potential, viscosity, and the
`clathric tendency (surface tension). The first two qualities
`are related to protein polarity; the second two relate to the
`diffusion of the protein, its partial molar volume, and to
`hydrophobic hydration.
`Osmolytic stabilizers. The first group of compounds are
`osmolytes, which are not strongly charged and have little
`effect on enzyme activity up to at least 1 M concentra—
`tion“. Their major effects are on the viscosity and surface
`tension of water, and hence on solvent ordering. Many of
`these compounds are used in vivo to control the osmotic
`pressure of eucaryotic and bacterial cells.
`Osmolytes can be polyols, sugars, polysaccharides, neu-
`tral polymers, amino acids and their derivatives, and large
`dipolar molecules like trimethylamine N-oxide (TMAO).
`Glycerol is the most commonly used osmolyte, as it is easily
`removed by dialysis and does not interfere with ion-
`exchange chromatography. It does not alter the dielectric
`constant of the medium significantly and its stabilization
`effect on proteins seems to be due to its ability to enter
`into and strengthen the water lattice structure. High
`concentrations of glycerol decrease the diffusivity and the
`partial molar volume of proteins”, thus lowering the rate
`of aggregate producing solute interactions.
`Glycerol has major drawbacks, however, especially for
`large scale work, as it is an excellent substrate for bacteria.
`Xylitol, a potential substitute,
`is not degraded well by
`bacteria and can be recycled from buffers by alcohol
`precipitation. PEG can be added to in vitro systems for
`nucleic acid and protein synthesis, where sufficient molec-
`ular density but low ionic strength is needed.
`Ionic stabilizers. Ionic compounds and salts can stabilize
`protein structure by shielding surface charges. Salts can
`also be considered as osmolytes and are used to some
`extent as such in vivo. E. coli transiently accumulated K+
`
`BIO/TECHNOLOGY VOL8 APRIL 1990
`
`and glutamate after osmotic shock, but within 30 min had
`switched to carbohydrates as osmoprotectantsi’. Most ionic
`compounds will affect
`the dielectric constant and the
`chemical potential of the solvent and the protein at
`concentrations well below where they affect the other bulk
`properties of the solution. Normal bacterial and mamma-
`lian enzymes function at a rather low salt concentration
`and are inhibited by high salt. Halophilic organisms,
`which can accumulate as much as 7 molal K+ intracellular-
`ly, have adapted their enzymes to function in very high
`salt concentrations“.
`
`There is no general rule on salting in of proteins;
`models that work for one protein are not necessarily
`applicable to another”. The salt concentration for maxi-
`mum solubility frequently falls within a very narrow
`range. As shown in Figure l, a 50 mM change in salt
`concentration gave as much as a 20-fold increase in
`dissolved T7 RNA polymerase. The solubilizing effect of
`ions are dependent on the size and charge distribution,
`but because polar groups on proteins are so diverse, it is
`hard to say a prion which ion will be best. Large ions are
`generally better at stabilizing proteins than small ones; in
`general, the more electronegative the ion, the more it
`interacts with and destabilizes protein structure.
`The finest experimental work on the effect of salts on
`protein solubility (usually during salting out) has been
`done by crystallographer52055v35. The assumed mecha-
`nism for salting out by small molecules is
`that
`they
`compete for water molecules until the concentration is too
`low to maintain the hydration sheath around the pro-
`tein“.
`
`Divalent cations can have extremely pronounced sol-
`ubility effects at very low concentrations. Even 1 mM Ca2+
`induces a conformational change characterized by insensi-
`tivity to trypsin in sea urchin hyalin, and Ca2+ and Mg2+
`in the range of 1—20 mM encourage self-association?
`Zn2+ aids in insulin solubilization as well as crystalliza-
`tion‘. As even tiny amounts of Cu, Zn, and Mn (among
`others) can also induce aggregation, chelators are often
`added to buffers.
`
`Denaturants, chaotrophs, myoprotectants, and other additives.
`One can solubilize almost any protein (usually at
`the
`expense of its activity) by chemical denaturation with
`perturbing ions. Urea stabilizes the unfolded states of
`proteins because essentially all protein parts, from the
`backbone to the tryptophan side chains, are more soluble
`in 6 M urea than water as evidenced by the free energy of
`transfer into this solvent”. Another class of denaturants,
`“chaotrophs” like guanidinium, cetyltrimethyl ammonium
`salts, trichloroacetate, and thiocyanate ions disrupt hydro-
`gen bond formation and disturb the hydration shell
`around proteins”. Detergents, amphiphilic compounds
`that lower the surface tension of water, bind to hydropho—
`bic areas of proteins.
`Another class of denaturants, organic solvents, lower
`the dielectric constant of water. The denaturing activity of
`hydrophobic solvents is due to a limited detergent effect
`and that they provide a competing interaction for the
`intramolecular hydrophobic interactions responsible for a
`stable tertiary structure. Some proteins are remarkably
`resistant to the denaturing effects of protic, hydrophilic
`organic solvents. The original method for isolation of
`insulin and human interferon-a from tissue and bacteria
`used extraction with acidic ethanol“; crambin can be
`crystallized from 60% ethanol”.
`Two organic solvents frequently used as cryoprotec-
`tants, dimethylsulfoxide (DMSO), and ethyleneglycol, can
`also denature proteins. DMSO encourages unfolding by
`favoring peptide N—H---O=S solvent bonds over peptide
`N—H---O=C peptide bonds“). Ethylene glycol, by reduc-
`
`
`
`Ex. 1022 - Page 3 of 10
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`

`

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`
`
`
`ing solvent polarity, weakens structural hydrophobic in-
`teractions.
`
`is general
`it
`Unless a protein is to be used in viva,
`practice to include protease inhibitors, sulfhydryl reduc-
`tants, bacteriocides, and chelating agents in small amounts
`to all protein solvents. The more common additives are
`listed in Table lc.
`
`Concentrating proteins. Limits on the maximum pro-
`tein concentration one can achieve are the structure of the
`protein,
`the buffer components and the purity of the
`protein preparation. Overloading the preparation on SDS
`acrylamide gels may not detect proteases that cause dam-
`age during concentration or storage. To minimize this
`contamination, during purification from bacterial ex-
`tracts, the protein should completely change buffer at
`least three times. Suitable transfer methods are salt pre-
`cipitation and dissolving in fresh buffer. binding to an
`affinity resin or HPLC column and elution, or gel filtra—
`tion. Dialysis, flow-through affinity steps, and redissolving
`lyophilized samples do not count as buffer transfers. All
`purification buffers should be made with ultra-pure water
`and HPLC grade chemicals where possible, and sterilized
`to avoid the reintroduction of bacterial contaminants.
`
`The most commonly used methods for concentration
`are salt precipitation, affinity chromatography, ultrafiltra-
`tion, and occasionally, chromatofocusing, electrofocusing,
`and freeze condensation (for cryoresistant proteins). Very
`stable proteins and peptides can be lyophilized or spray
`dried and redissolved. One should get the preparation to
`as high a concentration as possible by judicious elution of
`the last affinity step.
`The easiest method for concentrating proteins that
`cannot be lyophilized is ultrafiltration. Microconcentra-
`tors (Fig. 1) are useful for volumes up to 10 ml. Stirred
`pressure cells (Amicon, Millipore, or equivalent) are avail-
`able for volumes between 10 and 500 ml, and membrane
`type can be selected according to the size and hydropho-
`bicity of the protein. I was unable to use pressure cells for
`Mx protein or T7RP, however, as aggregation at
`the
`membrane surface was too high. The stir rate should be
`kept to a minimum as concentrated protein solutions are
`shear sensitive. For T7RP,
`losses were lowest with the
`Sartorius vacuum dialysis system, where I was able to
`concentrate to 40—50 mg/ml in 0.2 M ammonium sulfate
`buffer, pH 7.
`Hollow fibers or parallel plate continuous flow systems
`can be scaled up to any size. The Minitan system from
`Millipore is a good intermediate size for lab use. Protein
`loss on the membranes is significantly higher than the
`maximum predicted by the manufacturers.
`
`SITUATIONS WHERE PROTEIN SOLUBILITY
`BECOMES LIMITING
`
`Refolding inclusion body (18) proteins. 185 behave
`like protein that has been irreversibly precipitated. To
`obtain active protein, high concentrations of chaotrophic
`agents in the presence of sulfhydryl reducing agents are
`used to unfold the chains, which must then be refolded
`during removal of the denaturants. The primary refold-
`ing problem is aggregation of partially unfolded protein.
`In one study, the maximum protein concentration for
`efficient refolding was only 20 ug/ml‘“; for interleukin-2
`the maximum was only 1 pig/ml”. Concentration by
`ultrafiltration after refolding is possible, but losses due to
`proteolysis, aggregation of isomers, and membrane bind-
`ing are frequently very high. For tissue plasminogen
`activator (t-PA), the folding to intermediate states is rapid
`but the proper di—sulfide bonds form much more slowly.
`As the close to native folded forms are relatively soluble,
`timed addition of more unfolded protein concentrate (a
`
`sort of “fed batch") can allow much higher final concen-
`tration of the extract“? Residual denaturant can also
`stabilize the native state of the protein; its optimal concen-
`tration in the final extract should also be determined.
`
`TMAO may be a useful osmolyte when refolding proteins
`from urea solution“.
`
`Every protein contaminant present during refolding
`increases the total dilution necessary to avoid aggregation.
`In addition, partially unfolded proteins are excellent
`protease substrates. Thus one of the major advances in
`inclusion body protein refolding has been the develop-
`ment of purification steps that can be used in the presence
`of the denaturant. These include gel filtration, certain
`types of affinity chromatography”, and a new method
`based on the interaction between a poly-histidine peptide
`fused to the protein of interest and a nickel chelate
`column“.
`
`Alternate methods for refolding, such as binding dena-
`tured protein to thiol-sepharose columns or other affinity
`matrices and eluting with denaturant free buffers“, are
`also being explored. It
`is possible that activated thiol
`sepharose mimics the structure of protein disulfide isom-
`erases“. Serine proteinases46 and interleukin-4“7 refolding
`yields were greatly improved by pretreatment with gluta-
`thione. Interleukin-2 was renatured by dilution and auto-
`oxidation in the presence of Gun”.
`Appropriate choice of buffer during the refolding step
`can also improve yields at higher concentrations of pro-
`tein“. As optimal refolding conditions vary with the
`protein, one should either dilute the denatured sample
`into or dialyze it against many different buffers, and
`measure active or soluble protein after centrifugation.
`Solubilization and reconstitution of membrane en-
`
`zyme systems. Difficulties in solubilizing proteins from
`membranes have greatly limited structure and function
`studies“. Membrane proteins function in an amphiphilic
`environment and fold differently from cytoplasmic pro-
`teins: they turn their hydrophobic sites outward rather
`than inward. This probably accounts for why computer
`programs developed from soluble proteins predict the
`opposite of the known X-ray structures for membrane
`proteins”. This structural difference also accounts for the
`failure of detergents to solubilize IB proteins.
`The only way to isolate most integral membrane pro-
`teins is to extract them from their lipid environment with
`bulky detergents (typically Triton X-100 or Emulphogen
`BC-720). The protein is integrated into a detergent mi-
`celle with detergent replacing phospholipids or proteins
`that were previously in contact with the hydrophobic
`surfaces”. Even if the protein is not inactivated by this
`treatment, low critical micelle concentration (CMC) deter-
`gents interfere with protein concentration (by giving a
`gel), functional assays, and further purification steps (as
`the detergent’s properties dominate the protein’s).
`Thus proteins are transferred after the initial extraction
`to less harsh detergents forming smaller micelles18 via gel
`filtration. For detergents with CMC’s too low to allow for
`efficient dilution into monomers, one may need to use
`highly polar micelle dispersing agents like ethanediol or
`bile salts“). NMR structural studies of small membrane
`
`proteins in micelles“ is possible.
`A major advance in membrane protein crystallization is
`the use of “small amphiphiles” to replace detergents
`binding to the face of the protein“. One can thus prevent
`some of the problems caused by phase separation at
`higher salt and protein precipitation as the detergent in
`the micelles becomes too concentrated.
`
`Osmolytic stabilizers (20% glycerol) or high salt (0.3—0.4
`M KCl) added before the detergent may stabilize the
`tertiary structure of the protein during extraction and
`
`
`
`BIO/TECHNOLOGY VOLS APRIL 1990
`
`3“
`
`Ex. 1022 - Page 4 of 10
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`

`

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`dilution of the protein into proteoliposomes“. Glycerol or
`PEG is needed for efficient elution of membrane proteins
`from chromatofocusing columns“. In vitro assays of trans-
`port systems from bacteria24-52, signal peptidase from
`yeast“, and the tamoxifen binding protein from a breast
`cancer cell line55 were only possible byjudicious control of
`the salt concentration during detergent extraction of the
`membrane.
`
`As there is some evidence that high salt concentrations
`can stabilize secondary structural elements even during
`tertiary structure disruption“,
`the need for osmolytic
`stabilization may indicate that membrane proteins can
`undergo a transition phase, “molten globule” state during
`solubilization. This state is defined for soluble proteins as
`an intermediate during reversible unfolding which retains
`compact structure and CD spectrum similar to the native
`state, but shows other evidence (eg, increased binding ofa
`hydrophobic dye) of a non-native tertiary structure.
`Very concentrated protein solutions and NMR work.
`As growth factors and enzymes are so active, one general-
`ly works with solutions containing less than 1 lag/ml. But
`much more concentrated solutions are required for mi-
`croinjection into cells, for clinical trials of drugs and for
`analytical studies of protein structure. There are many
`references on preparing proteins for X-ray studies”? As
`it has only recently been shown to be a general method for
`protein structure determination“, less has been written
`on preparing proteins for NMR. The major requirement
`for good spectra is absolutely pure protein at high (1—20
`mM) concentration.
`Most structure determination by 1H-NMR used solu-
`tions in D20 and H20 at acid pH. Acid conditions
`encourage aggregation and protein unfolding, which
`shortens sample life. Solvent protons can significantly
`obscure regions of interest in the protein spectrum (Cu
`protons), so buffers are usually phosphate or deuterated
`Tris. Some groups prefer to work without ionic stabilizers,
`as they can blur peak profiles and cause excessive heat-up
`of the sample during measurements. These stringent
`requirements obviously limit
`the proteins that can be
`studied by the technique to small, stable ones.
`Assuming the solubility requirements are met, structure
`analysis for up to 80 amino acid proteins is almost
`routine“. The recent descriptions of well resolved (but
`very complex) 2—D NOESY and COSY spectra for uroki-
`nase (54 kD; solution was 1.5 mM in D20 at pH 4.5)“, as
`well as the interaction of pepsin (35 kD) with its ISN-
`labeled inhibitor59 show that investigation of even larger
`proteins is possible. Isotope-edited NMR spectroscopy,
`which selectively detects only protons bound to isotopical-
`ly labeled ('SN, 13C) nuclei, allows larger proteins to be
`analyzed and widens the choice of non-interfering buff-
`erSSS‘E’Q—SI.
`
`The solution should be stable during the measurement,
`which for larger proteins means addition of some salt.
`Staphylococcal nuclease was solubilized in 0.3 M NaCl at
`pH 7.6, ovomucoid domains (55 amino acids) were soluble
`to 12—15 mM in 0.2 M KCl at pH 86?, and yeast phospho-
`glycerate kinase substrate binding was studied in 0.1 M Na
`H—acetate buffer at pH 7.1 (unspecified enzyme concen—
`tration)“. Narrowest
`line widths were obtained for a
`solution of thrombin (35 kD) concentrated to 0.5 mM in
`0.2 M KCl at neutral pH. Significant line broadening was
`seen if the protein concentration was increased or at lower
`salt concentrations at the same pH (Gerhard Wagner,
`personal communication).
`
`PROTEIN ENGINEERING TO INCREASE
`SOLUBILITY
`
`Amino acid solubility and water aflinity. Individual
`
`amino acids vary greatly in solubility and affinity for water
`(Table 2). Protein solubility is based on the ability of
`soluble, polar residues to interact with water in such a way
`that the rest of the protein can maintain an active struc-
`ture. According to the “hydrophobic collapse" model of
`protein folding, the driving force for folding is hydropho-
`bic amino acid clustering to avoid water, with the eventual
`secondary and tertiary structure further stabilized by
`hydrogen bonding and electrostatic interactions”. The
`distribution of polarity toward the surface is so typical that
`it has been used as a criterion for protein design“.
`The data in Table 2 shows that the tendency of residues
`to be “buried” (definitions range from less than 5% of the
`residue surface exposed to solvent65 to up to 30%9) in a
`protein agrees with these generalizations. Most positively
`charged and amide side chain residues (His, Lys, Arg,
`Gln, Asn) were on the surfaces of the proteins studied,
`and the interiors were primarily composed of the alipha-
`tics Gly, Ala, Ile, Leu, Val and the aromatic Phe. But only
`23% of the Trp residues and 13% of the Tyr i