Refolding experiments after inclusion body proteolysis
Refolding experiments after inclusion body proteolysis
When proteins are expressed at high levels in E. coli and other expression hosts, it is common to find that most of them are insoluble and exist in insoluble forms called inclusion bodies. Although the precise mechanism of inclusion body formation is not fully understood and may vary depending on the protein and expression conditions, it is generally thought to be due to the fact that the rate of expression of the target protein is faster than the rate at which it folds out of its natural structure.
Authors: Burgess et al., Translated by Vivian Chen, this experiment is from "Protein Purification Guide".
Operation method
Refolding experiments after inclusion body proteolysis Move I. Routine operational programs This typical procedure below works well for many proteins. This operating scheme is adapted from one first developed by N g u y e n et al. (1993) and used in the insoluble recombinant protein purification section of the Cold Spring Harbor Protein Purification and Identification course (Burgess and K n u t h , 1996). Other similar processes may give similar results, but our laboratory uses this process almost exclusively. We will discuss the key steps below. (1) Using 2 L culture flasks containing IL L B medium, culture E. coli B L 21(D E 3)p L y S strain (Studieretal., 1"0) carrying the p E T vector (cloned with the target gene) at 37°C with shaking, straight A600nm to 0.6. The strain is then incubated with the p E T vector (cloned with the target gene) at 37°C with shaking. (2) Add isopropyl-β-D-galactosulfide (I P T G ) to 0.5~I m m o l /L to induce the expression of T 7 R N A polymerase, which then transcribes the target gene. (3) After 3~4 h of induction, cells were harvested by centrifugation at 15,000 r/min for 15 min. Cells were resuspended with a small volume of culture supernatant and then transferred to pre-weighed 40 mL Oak Ridge centrifuge tubes, and centrifuged to precipitate the cells. The wet weight of the precipitated cells is recorded and frozen at 80°C until ready for use. This method generally yields 1.5 to 2.0 g wet weight of E. coli cells per liter. (4) Thaw the cells, resuspend them in 30 mL of lysis buffer [50 mmol/L Tris-HCl (p H 7.9), 0.1 mmol/L LEDTA, 5% glycerol, 0.1 mmol/L DTT, 0.1 mol/L NaCl], and then ultrasonicate at 60% power three or four times for 20 s at intervals of 1 minute to cool on ice. (5) Add pure Triton X-100 (from 10 % by mass volume of stock solution) to 1 % to break down cell membranes and lyse membrane proteins. Incubate the lysate on ice for 10 m i n , then centrifuge at 15,000 rAni n for 15 m i n to precipitate inclusion bodies and remove the soluble supernatant. (6) Resuspend the inclusion bodies with 30 m L of lysis buffer containing 1 % Triton X-100, incubate for 10 m i n on ice, and then centrifuge for 15 m i n at 15 000 r /m i n . (7) The supernatant-removed inclusion body precipitate was resuspended in 30 mL of Trinton X-100-free lysis buffer to remove Trinton X-100, and then centrifuged as above. The resulting precipitate is referred to as the washed inclusion body fraction and is usually purer than 90 %. (8) The washed inclusion body precipitate is resuspended in a suitable denaturant and incubated to denature and solubilize, then centrifuged at 15,000 r/m i n for 15 m i n to remove any residual insoluble material. We usually use the above lysis buffer (without N a C l if diluted with guanidine hydrochloride) with the addition of 6 m o l/L guanidine hydrochloride (G u H C D, 8 m o l/L urea) or 0.3 % dodecylsuccinimide. 3 % sodium dodecyl sarcosinate [or N-lauroyl sarcosinate (sarkosyl or n-lauroyl sarcosinate)] was added to dissolve the inclusion bodies. (9) Recently, a relatively simple refolding assay (see below) was employed to identify suitable refolding buffers. (10) The protein concentration in the denatured sample was adjusted to I m g /m L . (11) Dilute the denatured protein 15 to 60 times to reduce the concentration of denaturant to a level where the protein can be refolded. Usually, we either dilute rapidly or slowly add the denatured inclusion body protein dropwise to the refolding buffer in a beaker with vigorous stirring for rapid mixing. The dilution is done at room temperature, and after mixing, the solution is allowed to stand for 1-2 h to ensure that the refolding process is complete and that aggregates are formed and flocculated. (12) Filter the refolded protein solution through a protein low affinity 0.22um membrane (e.g., Stericup--GV 0.22 m m o l /L, 500 m L, Millipore # SCG VU 05R E ) to remove particulate matter (if there are significant deposits, which may clog the membrane, filter the solution prior to filtration). the filter membrane, centrifugation may be performed prior to filtration). (13) Load the filtered solution into a 10 to 15 mL ion exchange column as quickly as possible (within column and system pressure limits, hopefully at least 10 mL/min). If conditions permit, monitor absorbance at 280 nm, 260 nm, and 320 nm. The absorbance at 320 nm is used to measure the scattering of light, and for some proteins, it shows the peak of multimer formation. (14) A 5 to 10 column volumes of buffer A [50 m m o l / L Tris-HCL (pH 7 . 9 ), 5 % glycerol, and 5 % glycerol] spiked with 0-I m o l / L NACL were used. 9 ), 5 % glycerol, 0.1 mol/L EDTA, 0.1 mol/L NACL, 0.1 mol/L EDTA, 0.5 mol/L NACL. The column was washed with [50 m m o l / L Tris-HCL (pH 7.9), 5 % glycerol, 0.1 mol/L EDTA, 0.1 mmol/L DTT], and then eluted with 10 column volumes of a linear gradient of 0.1-1.0 m o l / L NACL in the same buffer, at a flow rate of 5 m L / m i n , with 3-4: m L collected per sample. (15) Determine the purity of the different collection fractions by analyzing the 280 n m absorption peaks with S Ds- P A G E . If enzyme activity analysis is available, analyze each collection fraction to determine the fraction with the highest specific activity. (16) Mix the collection peaks, dialyze in lysis buffer containing 50 % glycerol, and freeze at 1 20 °C or 1 80 °C. (17) Characterize the mixed peaks. If possible, the specific activity is compared with that of a standard sample of the protein, which was prepared using more conventional methods not involving refolding. Obtaining high levels of expression is a different subject from refolding and will not be discussed further here (see Chapter 12 in this section ). Many systems can obtain expression levels of target proteins that account for 1 0 % ~4 0 % of the total cellular protein (Mak r i d e s , 1996; M u r b y et al. 1996; Sorensen and M o r t e n s e n , 2005; Studier et al,,, 1990) (see Section 9 in this chapter). Often, researchers need to perform experiments to determine at what temperature the cells should be cultured, how much inducer should be used, and for how long. When the target protein has been induced at a high level, the cells are usually centrifuged and then frozen at 180°C for later use. Some say that freezing the cells for weeks makes it difficult to refold the inclusion bodies, but in my experience, we can use cells that have been frozen for several months to do a good job of refolding. Unfortunately, however, as far as I know. We initially washed the inclusion bodies with 2 % sodium deoxycholate (Burgess andKrmth, 1996; Nguyenetal . , 1993), but later found that 1% Triton X-100 was more effective and easier to use. The use of Triton X-100 [Pierce, Surfact Amps X-100, 10 % (m/V) solution], which I call "Gourmet" (g o u r m e t), is highly recommended, as it has been purified to remove the peroxides found in Triton X-IOO, which is sometimes found in old yellowish bottles, and then stored in a nitrogen atmosphere in a sealed ampoule (amphora). o u l e) in nitrogen gas. The action of different salts and detergents can be experimented with to see which can more effectively wash out impurities from the inclusion bodies without solubilizing the target proteins, and sometimes 1 to 2 m d /L of urea can be used for washing. In general, it is necessary to wash out as much of the membrane components, D N A, and other proteins as possible before solubilizing the inclusion bodies. As stated previously, some inclusion proteins are near-natural and can be solubilized with mild conditions such as nonionic detergent or even 0-5 m o l / L N a C l (V e r a et al., 2006). In most cases, this will not work, and instead more intense denaturing conditions need to be used. As with the previous lysis buffer, a solubilizing agent/denaturant is typically dissolved in the buffer to control p H, chelate heavy metals, and maintain a reducing environment. Typically, the inclusion bodies are resuspended with the solubilizing agent, incubated for 30-60 min, and then centrifuged to remove insoluble material. Because the proteins in the inclusion bodies are at least partially denatured, dissolution can be done at room temperature, and sometimes it is even necessary to heat the solution to achieve complete dissolution. For example, when putting natural The following is a review of the use of several solubilizers. (1) Guanidine hydrochloride It is probably the most commonly used solubilizing agent, and its usual concentration in compatible buffers is 6 m o l /L. Most proteins are rapidly denatured in this strong ionizing agent (chaotropicagent), but in many cases incubation at higher temperatures will help to achieve complete denaturation of the protein. Once solubilized, dilution to 3 m o l /L guanidine hydrochloride is usually not a problem, since the change from the denatured state to the natural state usually occurs at a level of 1 to 2 m o l /L guanidine hydrochloride (P a c e , 1986). A dilution of ~0.1 m o l /L guanidine hydrochloride is usually sufficient to achieve a low enough salt level to ensure that the refolded target protein binds to the ion exchange chromatography column (see below). (2) Urea 8 m o l / L Urea is commonly used as a pro-solvent, especially for further purification of target proteins in the denatured state ( K r m t h and Burgess, 1987). In general, 8 m o l / L urea is not as effective as 6 m o l / L guanidine hydrochloride in promoting proteolysis and complete denaturation of proteins. It is important to note that cyanate, which is always present in urea solutions, may cause carbamylation of proteins (Chapter 44 of this book provides suggestions on how to minimize cyanate in urea). (3) Sodium Dodecyl Sarcosinate or Sodium Dodecyl Sulfate (sarkosyl or S D S ) Researchers have found that sodium dodecyl sarcosinate acts as an effective solubilizer to aid protein refolding at higher concentrations (Burgess, 1996). Sodium dodecyl sarcosinate is a very strong anionic denaturant, much like SDS, but it binds more weakly to proteins and dissociates more readily than SDS. Inclusion bodies are usually soluble in 0.3% sodium dodecyl sarcosinate, but it is important to note that sodium dodecyl sarcosinate should not be used to solubilize proteins equal to or greater than their own weight. For example, if you want to dissolve 150 m g of target protein in washed inclusion bodies, 20 m L of 0.3 % Sodium Dodecyl Sarcosinate (60 m g of Sodium Dodecyl Sarcosinate) will not completely dissolve the protein. You must add at least 50 m L of 0-3 % or 30 m L of 0.5 % Sodium Dodecyl Sarcosinate. We note that if there is a lot of Triton X-100 in the inclusion bodies, then more Sodium Dodecyl Sarcosinate is needed to solubilize the target protein. This is almost certainly due to the fact that Triton X-100 can form large microclusters, and can take up some Sodium Dodecyl Sarcosinate to form mixed microclusters, which are not pro-solubilizing. If a target protein solubilized with 0.3% Sodium Dodecyl Sarcosinate is diluted to about 0-01% Sodium Dodecyl Sarcosinate, most of the Sodium Dodecyl Sarcosinate is dissociated from the protein, and the protein is refolded. The residual detergent appears to bind to the hydrophobic regions (sticky sites) of the partially refolded protein, thus preventing aggregation. It acts like a chemical chaperone. If your target protein can bind to a cation-exchange column such as a POROSH, then the diluted solution can be filtered and pumped through a 10 mL column, washed with 10 to 20 volumes of 0.1 m o l/L NACL buffer, and eluted with a salt concentration gradient (see Section 5.5 of this chapter). The target protein will be bound to the column, while free Sodium Dodecyl Sarcosinate will flow through and residual Sodium Dodecyl Sarcosinate bound to the protein will be dissociated in the prolonged rinse. Experimental analysis of the eluted proteins showed that the proteins were essentially free (less than 1 molecule of sodium dodecyl sarcosinate in 10 protein molecules) of residual sodium dodecyl sarcosinate (R . B u rgess, unpublished data). Do not. SDS can be used in a similar manner, but care must be taken that the SDS used does not contain long alkanes (alkanes), such as C14, C1 6, etc. The SDS used in the SDS columns can also be used in a similar manner. Refolding experiments with GEP denatured in SDS have shown good results (R. Burgess, N. Thompson, and R. Chumanov, unpublished data), and therefore the use of SDS should be considered as an effective agonist. Successful pro-兾glutination and refolding of proteins using the cationic denaturant cetyltrimethylammo-nium chloride (CTAC) has been reported in the literature (Puriet al., 1992). The pro-solubilization effect was achieved even with distilled water with very low salt concentration (S o n g , 2009). Assuming that you have performed refolding experiments as described below, the most basic way to dilute denaturant is to dilute the solubilized protein into a suitable refolding buffer. However, there are three methods for refolding dilution. (1) Reverse dilution: The refolding buffer is added to the denatured protein and mixed well with each addition. There are several successful examples of this method [e.g., Grigskov and Burgess (1983)]. However, if one thinks about it, it is clear that this dilution method will undoubtedly produce the highest protein concentration during the critical period when protein refolding begins. For example, if the protein is dissolved in 6 m o l /L guanidine hydrochloride at a concentration of I m g /m L, then if 2 to 5 volumes of refolding buffer (diluted to 1 to 2 m o l /L guanidine hydrochloride) are added, it is possible that the protein may undergo the refolding transition at a concentration of 330 to 160 ug/mL. (2) Flash dilution: The denatured protein is quickly added to the refolding buffer. For example, 10 mL of refolded protein in guanidine hydrochloride at 6 mOl/L is added to 590 mL of suitable refolding buffer in a single step for a 60-fold dilution, with rapid mixing. At the time of refolding, the protein concentration will be 16ug/m L, which is well below the level of the reverse dilution, resulting in less likelihood of protein aggregation. (3) Drip dilution - The denatured proteins are added to the refolding buffer very slowly, drop by drop, over a period of I h. In theory, this should be optimal. Theoretically, this should be the best method because proteins always refold at very low concentrations [see S i n g h and P a n d a (2005), Vallejo and Rinas (2004) for a review]. For example, if you drop 0.1 m L of the above solubilized protein into 590 m L of refolding buffer, then the protein concentration is only ○.16ug/m L. Because you are dropping a very small increase in the amount of sample, the protein always stays at a low concentration, and when the last drop is added, the protein concentration is 16ug/m L. However, if the denatured protein undergoes a drop of I h and the However, if the denatured proteins are effectively refolded to their natural state after I h of dropwise addition, and the half refolding time for the proteins to effectively refold to their natural state is only 1 to 3 m i n , then the concentration of the partially refolded mucins will always be low, and thus they will not aggregate. It can be tiresome to spend i h adding solubilized proteins drop by drop, but it is possible to use a peristaltic pump set to a very low flow rate to add the denatured proteins over the i h period. This is much more reproducible and significantly reduces the effort expended by the graduate student. After the proteins have been refolded and filtered, the final step of the high-resolution ion exchange chromatography column is used to accomplish the following five important tasks. (1) Concentration of protein (e.g. from 600 mL to 4~8 mL) (2) Removal of denaturant (in flow-through solution) (3) Removal of minor impurities (in the flow-through solution, or binding to the column is weaker or stronger than binding to the target protein) If a high-resolution anion-exchange column is used, such as p ○ R ○ s H Q or M ○ n ○ Q, any D N A contaminants will be tightly bound to the column, and the use of ○ - 6 ~ ○ - 9 mol/L NaCl can be eluted. If it is highly desirable to have no E. coli lipopolysaccharides in the final product, the column can be rinsed with isopropanol before starting elution with the salt concentration gradient to greatly reduce LPS. (4) Selection of homogeneous, active monomers If samples of natural proteins purified by non-refolding techniques are available, then it is possible to know what salt concentration will elute them on the same ion exchange column. If a refolded sample elutes with a major peak at the same salt concentration, then one can be partially confident that the protein has been correctly refolded, since misfolded molecules may (but not necessarily) elute slightly differently if the column used is of high resolution. (5) Removing soluble protein multimers Soluble multimers are often present in refolded samples (see Section 5.7 of this chapter). If you have chosen a suitable refolding solution, there will probably be very few multimers; otherwise, they will be a major component of the refolding product. These soluble N polymers have JV times more charge than the monomers. Because of their higher charge, they tend to bind more tightly to the ion exchange column and elute later. Removal of the polymers is an important step if the protein is to be crystallized for structural determination, since heterogeneous phase materials are unlikely to crystallize. In my experience, in most cases, the protein in the first peak eluted from the column will be a high quality, pure protein with complete biological activity. If your protein does not contain cysteine, this is not a problem. However, most proteins contain cysteine and many have disulfide bonds which are an important part of the three-dimensional structure. Since the cytoplasm of E. coli is a very reducing environment, most endogenous proteins are in a reduced state. If you add a reducing agent to the lysis buffer, such as 0.1 ~ I rmnolZL of D T T , you can usually keep the proteins in a reduced state, preventing them from forming unwanted or incorrect disulfide bonds during the early steps of refolding described above. However, as soon as you refold the target protein, if it is capable of forming a disulfide bond, you must at some stage allow reoxidation of the protein to form the disulfide bond. In proteins that contain cysteines, but generally do not contain disulfide bonds, the cysteines are not in the precise geometrical configuration required to form disulfide bonds [see A nthony et al. (2002)]. Thus, although there are often a large number of possible erroneous disulfide bonds, they do not form readily. The best strategy is to perform protein refolding in a redox buffer (redox buffer). This buffer (see below) contains a mixture of reducing and oxidizing agents, which allows "shuffling" of disulfides to occur [see G ilbert (1994)]. The 'shuffling' of disulfides consists of the repeated formation and reduction of disulfide bonds. If an incorrect disulfide bond is formed in a misfolded protein and cannot be reduced, the protein is frozen in the wrong conformation and cannot reach its natural state. If this disulfide bond is reduced, the protein continues to vary between semi-folded intermediate states until the correct structure is formed. If the correct bond is formed, it stabilizes the final natural protein. Even if it is occasionally reduced, the protein will be in a stable state, and by reoxidizing it can again form the correct Some of the classical methods for forming disulfide bonds are described below. (1) Air oxidation Exposure to air without reductant for several days. (2) Redox buffers such as reduced glutathione (G S H )/oxidized glutathione (G S S G ) (10/1, 3 m m o l / L G S H /0. 3 m m m o l / L G S S G ). There are a variety of redox pairs (redox pairs) in use, which include reduced glutathione (G S H ) and oxidized cysteine, dithiothreitol (D T T ) and glutathione. The molar ratio of the reduced form to the oxidized form is sometimes different in order to achieve optimal reoxidation of the disulfide. (3) Protein disulfide isomerase (P D I ) This enzyme catalyzes the "shuffling" of disulfides (Kersteen and Raines, 2003). Small molecule PDI analogs (mimics) can also be used (W o y c e c h o w s k y e t a l . , 1999). The most common problem I find when browsing papers involving refolding is that the researcher does not know if the protein he ends up with has been folded correctly, is monodisperse, and has complete biological activity. Enzymatic or biological analysis is common, but there are usually no standards. You can see statements such as, "Since the end product is active, this indicates that it has been folded correctly and is fully active". However, you can only assess how much activity the end product has by comparing it to a known standard with complete activity. Without comparison, you can only know that it is active, but you cannot determine whether the percentage of active protein is 100, 100, 0.01 or 0.01%. 1 % or 0.01 %. If possible, it is necessary to determine the specificity and activity of the final product from the refolding process. Another important determination is the size of the refolded protein. Whether it is a monomer (monodisperse) or contains a large number of soluble polymers [heterodisperse]. Crystallographers commonly use dynamiclight SCattering (DSL) to determine whether purified, refolded or unfolded proteins are monodisperse. Another method, developed by Mark Knuth of the Genomics Institute of the Novartis Research Foundation (GNF) in La Jolla, California, uses analytical size exclusion chromatography (ANSEC) to determine whether purified, refolded, or unfolded proteins are monopersistent. The final product was analyzed by analytical size exclusion chromatography (ANSEC). We It is not possible to determine activity or mono-dispersity by circular dichroism (CD) or Western Blot analysis. There are numerous examples of proteins with very similar or identical CD spectra to natural standards that give good results in immuno-trace analyses but are in fact not mono-dispersive or inactive. In the previous routine process and discussion, it was assumed that a suitable refolding solution for the target protein is known. However, this is the most essential and difficult part of designing an effective refolding solution. In the earlier papers on protein refolding, only one refolding solution was usually chosen, and it was either effective or ineffective. Those that worked were published, and those that did not were usually discarded. This resulted in the early success of refolding proteins, which were then used in a variety of different ways. Several commercially available products have been developed to assist researchers in identifying suitable conditions for protein refolding. More information on protein refold screening and related protocols can be found on the company's website below. A t h e n a e s QuickFold™ (15-solution kit), http: //w w w w .athenaes.c o m /QuickFoldProteinRefolding Kit; EMD/Novagen, s iF0LDl™ , iF0LD2™ and iFOLD3™ (96-solution kit), http:,//www. novagen. com. Pierce Biotechnology's ProMatrix™ ( 96 -solution kit c o m p o n e n t s ) , http://w w w .fishersci.c o m . As experience with this screening technology is gained and new assists are identified, it is likely that this list will grow and the screening technology will continue to improve. It is important to note that for those researchers who cannot afford these expensive kits, there is nothing to prevent them from designing their own test solutions and developing their own protein refold screening methods to meet their specific needs, the key here being the systematic parallel screening of multiple refolding conditions. There are many solution variables (e.g., pH, temperature, salt concentration, oxygenation environment, and presence of divalent ions) and additives (which have been reported to increase the refolding efficiency of dissolved inclusion proteins). These variables and additives have been taken into account in the design of the commercialized protein refolding screen products mentioned earlier, and they provide examples for researchers who wish to design their own protein refolding screen methods. These variables are discussed below. Some of these variables are discussed in more detail in other reviews of protein refolding (Armstrong et al. 1999; Coweson et al.> 2006; Middelberg 2002; Quronfleh et al.> 2007; Schein> 1991; Singh and Pattan, 2005). and P a n d a , 2005; Tresaugues et al. 2004; Vallejo and Rin a s , 2004; Vincentelli (1) The refolding of most proteins with p H is completed within 5 to 9 p H. In our experiments, most proteins are best refolded at 8 to 8.5 p H. In general, the p H used is close to the isoelectric point of the protein (the protein does not carry a net charge and is most inclined to the sinking point). In general, it is not a bad idea to use p H at least one unit away from the isoelectric point of the protein (the point at which the protein has no net charge and is most inclined to the p H at the time of precipitation). (2) Temperature At present, there is no clear trend towards any widely applicable optimal protein refolding temperature. Most researchers perform protein refolding at temperatures close to room temperature. This temperature is low enough to prevent thermal damage to proteins, but high enough to increase the thermodynamic motion of the molecules, which may be important for dissolving transient misfolded conformations and achieving the natural conformation. One might argue that higher temperatures enhance the hydrophobic interactions that can lead to aggregation, but it also enhances the hydrophobic interactions required to bury hydrophobic residues inside the natural conformation. Xie and Wetlaufer (1996) published a very interesting paper on the refolding of the enzyme IKcarbonic anhydrase II at different temperatures at a protein concentration of 4 m g / m L, I m m o l / L GuHCl and 50 m m m o l / L Tris sulfate, p H 7. 5. anhydrase II). The results showed that refolding at low temperatures (4~12°C) for 120 min, followed by a temperature "temperaturundefinedleap" to 36°C for 30 min, gave a very good recovery of enzyme activity (>90 %). They concluded that hydrophobic interactions are reduced at low temperatures, minimizing aggregation while allowing the protein to slowly transform into an inactive or non-aggregating intermediate. When the temperature 'jumps' from 4°C to 36°C, this intermediate is converted to a highly reactive natural form. (3) Salt concentration may require certain salts to act as "salting in" to increase the solubility of natural proteins (see Chapter 20 of this book). Typically, 50 to 100 m mol/L of salt is used. After a 60-fold dilution of 6 m mol/L hydroxide fox, the final concentration of guanidine hydrochloride becomes 0.1 m mol/L. To increase the solubility of natural proteins, it is necessary to dilute the salt by a factor of 0.1. The additio For more product details, please visit Aladdin Scientific website.
Unfortunately, to my knowledge, there is no systematic study that addresses this issue.
For example, when the natural green fluorescent protein (GFP) is dissolved in 6 m ol/L guanidine hydrochloride, it remains fluorescent at room temperature but denatures and loses its fluorescence up to Imiri at 75°C (R. Burgess and N. Thompson, unpublished results). A more detailed discussion of solubilizing effects can be found in Marston and Hartley (1990).
do this with a M o n o S column because sodium dodecyl sarcosinate will bind to the column and damage it, and it will contaminate your eluted proteins.
After the protein is diluted it is allowed to stand for 30 to 60 m in and then filtered as described in the routine procedure. Otherwise, when the protein solution is loaded onto the column (see below), very small particulate matter may clog the column, causing a pressure rise that can cause the experiment to fail, or the flow rate may need to be reduced so low that loading the sample takes hours.
Bonds are formed again by reoxidation. For this reason, the addition of reducing agents only to the lysis buffer, but not to the refolding buffer, is often successful. After refolding, proteins can be subjected to slow air oxidation (exposure to air for 1 to 2 days), which often results in the correct disulfide bond structure. More information on reoxygenation can be found in the literature of V allejo and Rinas (2004) and Kirsten and Raines (2003).
We have found this method to be very useful because even very small amounts of protein can be analyzed by this method. Typically, we analyze 20-50 ug of sample at a flow rate of 0.5 to 1.0 mL/min using a 12 mL Shedex KW-802.5 column and a buffer containing 0.25 mOl/L NaCl, while monitoring ultraviolet absorptions of 280 rnn and 215 nm. Once the column has been calibrated with an appropriate molecular mass standard, it can be quickly determined whether the bulk of the protein is within a single peak formed by monomers of the predicted size (unidispersity, good condition) or whether the bulk of the protein has been washed out earlier, which implies that polymer formation has resulted in a larger molecular mass (non-homogeneous dispersion, bad condition). This step of analysis is an important component of a complete refolding test (see below).
The result was that the proteins that were successfully refolded in the early stages were those that usually refolded readily under many different conditions. In recent years, it has become increasingly clear that many proteins can be refolded only under very specific conditions. The challenge has become how to sift through the many possible conditions to select those that effectively promote protein refolding. One of the most important advances has been the use of the fractional factorial approach to systematically determine the effects of many different variables (Armstrong et al., 2004).
Strong et al. 1999; Cowieson et al. 2006; Quronfleh et al. 2007; TTre saugues et al. 2004; Vincentelli et al. 2004; Willis et al. 2005).
et a l , 2004; Willis et al.)