Appropriate regeneration of StrepTrap HP columns for FPLC

Appropriate regeneration of StrepTrap HP columns for FPLC

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My question is related to protein purification using a ÄKTA FPLC. We used StrepTrap HP Columns (1 ml column Volume (CV)) from GE Healthcare Life Sciences to purify a strep-tagged protein. In the first approach, this worked pretty fine. We wanted to re-use the columns after regenerating them according to the official protocol. The included steps were all performed at a flow rate of 1 ml/min and with filtered solutions:

  1. 3 CV distilled water
  2. 3 CV 0.5 M NaOH
  3. 3 CV distilled water
  4. 5 CV 20 % EtOH
  5. Storage of columns in 20 % EtOH at 4°C.
  6. Equilibration of the columns with Binding Buffer just before next use.

By adhering to the protocol we encountered some problems with the next purification (it was the same protein, but a new sample of cell extract). The purification worked less effective and the amount of purified protein was very low.

We tried a slightly different protocol for regeneration, also at a flow rate of 1 ml/min and filtered solutions.

  1. 5 CV distilled water
  2. 5 CV 20 % EtOH
  3. 3 CV 6 M Guanidinhydrochloride (sol.)
  4. 5 CV 0.5 M NaOH
  5. 3 CV distilled water
  6. 5 CV 20 % EtOH
  7. Storage in 20 % EtOH at 4°C
  8. Equilibration with Binding Buffer before next use.

We chose the Guanidinhydrochloride to denaturate all remaining proteins from the column resin and support the regeneration by this. But also with this protocol we encountered the same problems (low efficiency of purification).

I would be very interested if anyone of you have encountered the same problems and maybe have a solution/better protocol for this regeneration step.

Thank you for your suggests in advance! :)

Structure determination protocol for transmembrane domain oligomers

The transmembrane (TM) anchors of cell surface proteins have been one of the ‘blind spots’ in structural biology because they are generally very hydrophobic, sometimes dynamic, and thus difficult targets for structural characterization. A plethora of examples show these membrane anchors are not merely anchors but can multimerize specifically to activate signaling receptors on the cell surface or to stabilize envelope proteins in viruses. Through a series of studies of the TM domains (TMDs) of immune receptors and viral membrane proteins, we have established a robust protocol for determining atomic-resolution structures of TM oligomers by NMR in bicelles that closely mimic a lipid bilayer. Our protocol overcomes hurdles typically encountered by structural biology techniques such as X-ray crystallography and cryo-electron microscopy (cryo-EM) when studying small TMDs. Here, we provide the details of the protocol, covering five major technical aspects: (i) a general method for producing isotopically labeled TM or membrane-proximal (MP) protein fragments that involves expression of the protein (which is fused to TrpLE) into inclusion bodies and releasing the target protein by cyanogen bromide (CNBr) cleavage (ii) determination of the oligomeric state of TMDs in bicelles (iii) detection of intermolecular contacts using nuclear Overhauser effect (NOE) experiments (iv) structure determination and (v) paramagnetic probe titration (PPT) to characterize the membrane partition of the TM oligomers. This protocol is broadly applicable for filling structural gaps of many type I/II membrane proteins. The procedures may take 3–6 months to complete, depending on the complexity and stability of the protein sample.


The R-Spondin (roof plate-specific Spondin) proteins comprise a family of secreted proteins, which are known for their important roles in cell proliferation, differentiation and death, by inducing the Wnt pathway [1, 2]. RSPOs are expressed in several embryonic tissues and in the adult, with adequate expression levels being essential for organism development and homeostasis maintenance [3, 4]. Several studies have demonstrated the importance of RSPOs in regulation of several tissue-specific processes, such as bone formation, skeletal muscle tissue development, pancreatic β-cells and intestinal stem cells proliferation and even cancer, as reviewed by Yoon and Lee [5]. However, inappropriate functioning of these proteins may lead to various pathological conditions [reviewed by [5, 6], such as: sexual phenotype reversal, hyperkeratosis and a predisposition for squamous cell carcinoma of the skin [7, 8], craniofacial defects and problems in limbs, lungs and hair follicles formation [9,10,11,12], in placental formation [13, 14] and in the development of nails (Anonychia) [15,16,17,18]. Therefore, it becomes evident that RSPOs display a great therapeutic potential for treatment of a number of diseases.

Four RSPOs proteins (RSPO1 through 4) have been described, all of which display characteristic domains which are conserved among vertebrates, such as: (1) a thrombospondin 1 repeat (TSR) domain, (2) a cysteine-rich furin-like (CR-FU) domain, (3) a basic amino acid-rich (BR) domain of variable length (carboxy-terminal region), and (4) a hydrophylic signal peptide (SP) sequence [5]. The signal peptide present at the protein amino-terminal region ensures its entry into the canonical secretory pathway, addressing to the endoplasmic reticulum, transit through the Golgi apparatus complex and secretion to the extracellular space [4, 19]. The RSPO CR-FU domain, in turn, is identified as being responsible for mediating the Wnt/β-catenin signaling pathway activation [4, 19,20,21], although other studies also suggest that this domain may be involved in regulating the secretion of these proteins [21]. On the other hand, the BR and TSR domains have been proposed to be responsible for regulating the intensity of RSPOs action in the canonical Wnt pathway induction [20]. In addition, the TSR and BR domains still seem to be responsible for the association of RSPOs to the extracellular matrix (ECM), by binding to glycosaminoglycans (GAG) and proteoglycans [4, 19, 22,23,24] [reviewed by [6]].

Currently, it is known that RSPO proteins are capable of inducing the canonical (beta-catenin-dependent) and non-canonical (beta-catenin-independent) Wnt pathways [reviewed by [5, 6]]. However, although several studies on RSPOs mechanisms of action are available, many questions still remain about their receptors and the mechanisms involved in signal transduction by these proteins. Studies revealed that RSPOs bind to leucine rich repeat containing G protein-coupled receptors 4–6 (Lgr4–6) [25,26,27] to induce the canonical Wnt/beta-catenin signaling pathway, but other studies also indicate that these proteins are able to bind to low-density lipoprotein receptor-related proteins 5–6 (Lrp5–6) [4, 19, 21, 28] and Kremen1 (KRM1) [29]. It is also known that RSPOs act by inhibiting the Zinc and Ring Finger 3 protein (ZNRF3) [30] and binding to Frizzled 8 (Fzd8) to induce the Wnt pathway [19], although, apparently, RSPOs only bind weakly to the FZD receptors [21, 28]. However, part of this controversy regarding RSPOs receptors may be explained by studies suggesting a synergistic action of these proteins with Wnt ligands [4, 31]. In addition, in a recent work, the non-equivalence of the WNT proteins and the RSPOs with respect to the induction of self-renewal in LGR5 + intestinal stem cells was observed, but cooperation between these proteins was highlighted [32]. Unlike the canonical Wnt pathway, the β-catenin-independent pathway seems to have a less contradictory status in the literature, although it is less explored and still presents several gaps. Recently, syndecans were found to be new RSPOs receptors in the Wnt pathway [24], but studies have shown that only RSPO2 and RSPO3 proteins bind to syndecans.

RSPO1 stands out among RSPOs molecules with respect to its potential therapeutic use, especially in the Regenerative Medicine field, due to its mitogenic activity in stem cells. This potential has been confirmed by several studies which have shown the use of RSPO1 in several animal models for treatment of: intestinal mucositis induced by chemotherapy [33] or radiation [34], inflammatory bowel diseases (IBD) such as ulcerative colitis (UC) and Crohn’s disease (CD), in which the inflammatory response leads to continuous intestinal epithelial cells death [31, 35] and Diabetes Mellitus (DM), as a cytoprotective and proliferative agent for β-cells, by regulating the canonical Wnt pathway [36, 37]. Furthermore, other studies suggest its use in joint diseases such as arthritis [38] and in cancer, possibly acting as a tumor suppressor gene in lymphocytic leukemia [38, 39].

The RSPO1 protein, produced and studied here, is comprised of 263 amino acid residues arranged in a single 28,959 Da polypeptide chain. According to the Universal Protein Resource database (UniProt), RSPO1 is encoded from the RSPO1 gene, located on chromosome 1, position 38,076,951 to 38,100,595, presenting four isoforms, which arise from alternative exons splicing. In silico analysis of the RSPO1 protein demonstrated a three-dimensional structure rich in β-sheet secondary structures, lacking alpha helix. Recent studies demonstrated the presence of an N-glycosylation at the asparagine Asn137 of the polypeptide chain, which is related with RSPO1 secretion, activity and stability [40, 41], although both articles mentioned above present conflicting results about the effect of N-glycosylation on the biological activity of RSPO1 protein.

Here, the recombinant human RSPO1 was generated using a human cell line, namely: HEK293 (Human Embryonic Kidney) cells. Mammalian cells have been used for the production of several recombinant proteins, especially due to their ability to carry out post-translational modifications, which are essential for maintaining the structure and function of proteins. Among the post-translational modifications, glycosylation deserves special attention in the production of recombinant proteins in heterologous systems, since these modifications can interfere with protein folding, activity, stability and maturation, depending on the expression system used [42]. In this context, due to its capacity to generate complex glycosylation patterns, especially with the addition of sialic acids, the HEK293 cell line has been widely used for the production of recombinant proteins, being the human cell line most often used in the production of biopharmaceuticals approved by regulatory agencies, such as the FDA (Food and Drug Administration) [43, 44].

The objective of this study was to generate a stable expression platform for production of rhRSPO1 in human cells in order to obtain a purified, characterized and biologically active protein product. In the future, this platform may be optimized for rhRSPO1 production in an efficient and reproducible manner to be used in cell therapy. In addition to generation of rhRSPO1 overproducing cell clones, a new rhRSPO1 purification protocol has been established yielding a high purity protein product.

Results and Discussion

There are two distinct aspects of the procedure introduced here. First, multiple components of the expression mixture were individually screened for optimized formation of native disulfide bonds. Second, it is shown that these optimized solution conditions can be combined with GB1 (the B1 domain of protein G from Streptococcus sp.) fusion for enhanced expression [ [21] ]. The success of this combination is linked to subtle details in preparing the reaction mixture.

Activity of cell-free protein synthesis under oxidizing conditions

The translation machinery in the starting expression system for this project [ [21] ] was derived from the reducing environment of the E. coli cytoplasm. We therefore first determined the protein synthesis yields in this system under increasingly oxidizing conditions. Small-scale expression of the membrane protein OmpX, which is exclusively produced in the insoluble fraction, was performed in reaction mixtures supplemented with variable ratios of reduced glutathione (GSH) and GSSG (Fig. 1). SDS/PAGE analysis of the insoluble expression product indicated that the intrinsic protein synthesis is not inhibited by addition of up to 10 m m GSSG, even in the absence of GSH (Fig. 1). At a level of 20 m m GSSG the production of OmpX decreased significantly, and at 30 m m GSSG the cell-free system no longer yielded measurable synthesis of OmpX. These results can be rationalized by the observation that protein components of the cell-free system precipitated at GSSG concentrations above 10 m m , and so they could be detected in the insoluble protein fraction. Addition of up to 20 m m GSH in the absence of GSSG did not measurably affect the translation efficiency.

Impact of redox conditions on the translational activity of the E. coli based cell-free expression system [ [21] ] that was used as the starting point for the present study. Cell-free expression of the test protein OmpX in pIVEX2.4 was conducted for 2 h at 30 °C in the presence of the indicated ratios of GSH and GSSG. After the reaction, the insoluble protein fraction containing OmpX was collected and analyzed by SDS/PAGE. Each lane corresponds to 3.5 μL of reaction mixture. N indicates the negative control reaction without template DNA.

Fine adjustment of redox conditions for disulfide bond formation in the cell-free system

To determine appropriate redox conditions that favor disulfide bond formation and native folding without sacrificing translation efficiency, we investigated soluble production of hDpl(24–152)-pIVEX2.4 in cell-free reaction media supplemented with various ratios of GSH and GSSG. Western blot analysis of the reaction supernatant revealed that the highest soluble expression of hDpl(24–152) was obtained in the presence of 2 m m GSH and 5–10 m m GSSG (Fig. 2A). Since the reaction mixture also contains 2.1 m m dithiothreitol (DTT), 2.1 m m β-mercaptoethanol (2-ME) and 1.5 m m cysteine (Table 1), high soluble expression was thus obtained in the presence of about 10 m m free sulfhydryl groups and 5–10 m m GSSG.

Influence of variable redox potentials and disulfide bond isomerase concentrations on the cell-free expression of three target proteins containing disulfide bonds. Shown are western blot (A, B, C) and SDS/PAGE (D) analyses of the soluble proteins obtained after cell-free expression during 2.5 h at 30 °C using either the pIVEX2.4 (A, B, D) or the pCFX1 (C) vector. For each lane in (A), (C) and (D) the applied sample volume corresponds to 2.5 μL of reaction mixture, and in (B) to 1.5 μL. (A) Soluble expression levels of hDpl(24–152) at various redox potentials achieved by mixing different ratios of GSH and GSSG. (B) Reaction mixtures for expression of hDpl(24–152) that contained 2 m m GSH and 5 m m GSSG were supplemented with increasing concentrations of the disulfide bond isomerases DsbC or PDI. (C) Soluble cell-free expression of GB1–mDpl(24–155) at variable levels of the redox potential and DsbC concentration. (D) Effects of variable redox potentials and DsbC concentrations on the soluble production of mIL22(34–179). Arrows indicate the bands corresponding to the target protein, and N identifies the lane of the negative control reaction without template plasmid.

Component Concentration
KOAc 217 m m
Creatine phosphate 80 m m
HEPES-KOH 58 m m
Mg(OAc)2 11 m m
NaN3 3.8 m m
PEG-8000 3.25% (w/v)
2-ME 2.1 m m
DTT 2.1 m m
ATP 1.2 m m
GTP 0.86 m m
CTP 0.86 m m
UTP 0.86 m m
cAMP·Na + 0.64 m m
Folinic acid·Ca 2+ 68 μ m
E. coli tRNA 175 μg·mL −1
Creatine kinase 5.8 μ m
Target plasmid 10 μg·mL −1
T7 RNA polymerase 65 n m
Amino acids (each) 1.5 m m
S30 cell extract 30% (v/v)

Supplementing the cell-free expression system with disulfide bond catalysts

SDS/PAGE comparison of both soluble and insoluble yields of hDpl(24–152) revealed that only about 5% (8 μg·mL −1 ) of the total hDpl produced (160 μg·mL −1 ) was obtained in soluble form at the aforementioned optimal redox conditions. In an attempt to increase soluble production of hDpl(24–152), the reaction mixture with redox conditions fixed at 2 m m GSH and 5 m m GSSG was supplemented with various concentrations of thiol : disulfide interchange protein DsbC (DsbC) and protein disulfide isomerase (PDI). Western blot analysis showed that a supplement of 10 μ m DsbC or PDI significantly increased soluble production of hDpl(24–152) (Fig. 2B). Thereby more than 50% (80 μg·mL −1 ) of the synthesized hDpl(24–152) was obtained in soluble form. The conditions thus established for optimal expression of hDpl(24–152) could be carried over to GB1–mDpl(24–155) (Fig. 2C).

For mIL22(34–179), SDS/PAGE analysis indicated that, in the absence of enzymatic disulfide bond catalysts, maximal production of soluble protein was achieved after addition of 2 m m GSH and 5–10 m m GSSG. Addition of 5 μ m DsbC further increased production of the soluble protein ∼ 2.5-fold (Fig. 2D), and more than 80% (0.2 mg·mL −1 ) of the total synthesized mIL22(34–179) (0.25 mg·mL −1 ) was obtained in soluble form. Since maximum soluble yields were obtained with 5 μ m DsbC, there is an indication that the native arrangement of disulfide bonds is more readily achieved for mIL22(34–179) than for hDpl(24–152) or for mDpl(24–155).

The disulfide bond oxidase thiol : disulfide interchange protein DsbA (DsbA) has previously been shown to enhance native folding of proteins in vitro by enhancing disulfide bond formation during the folding process [ [25] ]. We therefore also investigated the effects of adding up to 25 μ m DsbA, prepared as described in Doc. S3, to a reaction mixture that contained also 2 m m GSH, 5 m m GSSG and 10 μ m DsbC. Under these conditions the addition of DsbA did not increase the soluble yield of hDpl(24–152). Similarly, addition of DsbA in the absence of DsbC did not enhance soluble expression, and we did not further pursue experimenting with DsbA. Overall, both the prokaryotic DsbC and the eukaryotic PDI promoted expression of soluble target protein, indicating that they can be used interchangeably in the present expression system.

Cell-free expression with S30 cell extract from E. coli Origami (DE3) cells

E. coli strains containing mutations in the genes encoding thioredoxin reductase (trxB) and glutathione reductase (gor) have been shown to yield enhanced disulfide bond formation in the bacterial cytoplasm [ [26] ]. To investigate the potential of extracts from these strains for cell-free production of proteins containing disulfide bonds with the protocol optimized for BL21 (DE3) RIPL-Star, we prepared S30 extract from the E. coli Origami (DE3) strain (Novagen, Darmstadt, Germany). We then compared cell-free production of mIL22(34–179) using S30 extracts prepared from either E. coli Origami (DE3) or E. coli BL21 (DE3) RIPL-Star cells, which were supplemented with various ratios of GSH and GSSG. Analysis by SDS/PAGE indicated very low expression levels of mIL22(34–179) in reactions with S30 extract from Origami cells, and we did not further investigate this extract.

N-terminal GB1 fusion for increased expression yields

We recently showed that N-terminal fusion constructs with the GB1 domain have increased yields of cell-free expression of human proteins [ [21] ]. Here we explore the use of GB1 fusion with the aforementioned cell-free systems that have been optimized for disulfide-containing proteins. Overall, GB1 fusion of the target proteins was thus found to affect neither the optimal GSH/GSSG ratio nor the optimal concentration of disulfide bond isomerase required for native folding. Thus 10 mL cell-free reactions of hDpl(24–152) with (pCFX1) or without (pIVEX2.4) fusion with the GB1 solubility domain were carried out at optimum conditions for disulfide bond formation, i.e. with 2 m m GSH, 5 m m GSSG and 10 μ m DsbC. After purification, UV absorption at 280 nm indicated yields of 4.2 mg soluble hDpl(24–152) from pCFX1 and 0.8 mg from pIVEX2.4. The 5-fold increase for the fusion construct with the GB1 domain is probably due to both increased total production of the protein by enhanced translation efficiency [ [21] ] and increased solubility [ [27] ]. SDS/PAGE showed that the yield of 4.2 mg corresponds to 75% of the total hDpl(24–152) expression (5.6 mg), compared with ∼ 2% of the total synthesized protein in the absence of disulfide-forming additives.

In further assays, 10 mL reactions of mIL22(34–179) in either pIVEX2.4 or pCFX3 were carried out using [u- 15 N]-amino acids at optimum conditions for mIL22(34–179), i.e. with addition of 2 m m GSH, 10 m m GSSG and 5 μ m DsbC. We obtained 2.0 mg of mIL22(34–179) from expression in pIVEX2.4 and 6.6 mg from expression in pCFX3, where SDS/PAGE indicated that more than 90% (6.6 mg) of the total produced N-terminal fusion construct with the GB1 domain (7.3 mg) was soluble. In the absence of disulfide-promoting additives, only ∼ 5% of the expressed GB1–mIL22 fusion protein was soluble. Finally, a 10 mL cell-free reaction using [u- 15 N]-labeled amino acids in pCFX1 with addition of 2m m GSH, 5 m m GSSG and 10 μ m DsbC yielded 3.0 mg of purified mDpl(24–155), corresponding to ∼ 50% of the total expressed protein (6 mg). The lower yield of soluble protein, compared with hDpl(24–152), seems to reflect increased difficulty in producing the mouse protein, as was previously observed with expression in E. coli cell cultures [ [23, 24] ]. The yield of soluble GB1–mDpl fusion protein without disulfide-promoting additives was ∼ 2% of the total synthesized protein.

Structural validation of the disulfide-containing proteins from cell-free production and conclusions

The free sulfhydryl content of the three proteins studied here was determined with the Ellman assay under denaturing conditions, as described in the Materials and methods section. There was no evidence of measurable free sulfhydryl content, implying that the disulfide bonds had been formed successfully.

To obtain evidence that the proteins produced by cell-free expression were natively folded, we recorded 2D [ 15 N, 1 H] heteronuclear single quantum coherence (HSQC) NMR spectra. For cell-free produced hDpl(24–152) and mDpl(25–155), comparison with the available backbone amide resonance assignments [ [23, 24] ] indicated that the native fold was obtained (Fig. 3). This includes that hDpl(24–152) expressed either free or as a fusion construct with the GB1 domain adopted the native fold (Fig. 3A,B). The spectra also contain NMR signals from the N-terminal purification and solubility tags.

2D [ 15 N, 1 H]-HSQC spectra of hDpl(24–152) and mDpl(24–155) prepared by cell-free expression with 15 N-labeled amino acids in reaction mixtures containing 2 m m GSH, 7.5 m m GSSG and 10 μ m DsbC. (A) 200 μ m [u- 15 N]-hDpl(24–152) produced from two 10 mL reactions using the pIVEX2.4 vector. (B) 275 μ m [u- 15 N]-GB1–hDpl(24–152) prepared from a 10 mL cell-free reaction using the pCFX1 vector. The additional signals originating from the GB1 domain [ [27] ] can be readily recognized. (C) 140 μ m [u- 15 N]-mDpl(24–155) after thrombin cleavage to remove the N-terminal GB1 solubility domain. The protein was synthesized in a 10 mL batch-mode cell-free reaction using the pCFX1 vector. All samples contained 20 m m sodium acetate at pH 5.2, 100 μ m EDTA, 10 μ m sodium azide and 5% D2O. The spectra were recorded at a 1 H resonance frequency of 750 MHz with T = 20 °C. Red crosses in (A) and (B) show the published peak positions for hDpl(24–152) (BMRB 5145), and those in (C) the peak positions for mDpl(26–157) (BMRB 4938).

Human interleukin-22 had been shown to form dimers in solution even at micromolar concentrations [ [28] ]. It appears that mIL22(34–179) is also dimeric in solution, since the 2D [ 15 N, 1 H]-HSQC spectrum contains a lesser number of signals than expected from the number of amide groups in the protein, with most residues represented by weak peaks (Fig. 4A). It can nonetheless be recognized that the NMR spectrum corresponds to a folded globular protein. For the denatured protein in urea, the 2D [ 15 N, 1 H]-HSQC spectrum contained a number of amide resonances which, within the accuracy of the peak count, agreed closely with the number expected from the primary structure (Fig. 4B). The cell-free produced mIL22(34–179) showed a far-UV CD spectrum typical of a protein with a high content of helical secondary structure (Fig. 5), and biological activity of the produced protein was verified using IL-22-deficient mice in the laboratory of B. Becher [ [29] ].

2D [ 15 N, 1 H]-HSQC spectra for (A) native and (B) urea-unfolded [u- 15 N]-mIL22(34–179) expressed with the vector pIVEX2.4 in a 10 mL cell-free reaction mixture supplemented with 2 m m GSH, 10 m m GSSG and 5 μ m DsbC. In (B) the protein was denatured in 8 m urea and 10 m m DTT. The spectra were recorded at a 1 H resonance frequency of 750 MHz and at T = 37 °C.

Far-UV CD spectrum at 20 °C of mIL22(34–179) produced by cell-free synthesis. The sample contained 15 μ m mIL22(34–179), 3 m m sodium phosphate at pH 7.0, 34 m m sodium chloride and 1.1 m m potassium chloride.

Overall, the present documentation of native folding of disulfide-containing eukaryotic proteins produced in milligram quantities in a cell-free system validates the cell-free expression protocol used, which included adjusting an appropriate redox potential by addition of small concentrations of GSH and GSSG to S30 cell extract from the E. coli BL21 (DE3) RIPL-Star strain and supplementing this mixture with a disulfide bond isomerase to facilitate the correct arrangement of disulfide bonds. It is worth noting that the best results were obtained in the presence of 10 m m sulfhydryl groups and 5–10 m m GSSG, which corresponds closely to the redox conditions in the endoplasmatic reticulum [ [30] ], where disulfide bonds are typically formed [ [2] ]. The renewed observation of significantly increased soluble expression of N-terminal fusion constructs of the target protein with the GB1 domain [ [21] ] indicates that this approach might be quite widely applicable to obtain increased yields in cell-free expression. With regard to using this expression system in structural biology, it is of interest that energy regeneration is based on creatine phosphate, which does not activate metabolic pathways in the cell extract and therefore enables the production of stable-isotope-labeled proteins. It further avoids reduction of GSSG during the cell-free reaction and thus eliminates the requirement for chemical pretreatment of the S30 extract. These results set our expression system apart from previously described cell-free systems. These either provided yields of < 50 μg native protein per milliliter of reaction mixture [ [9-11, 14] ] or were based on energy regeneration with phosphoenolpyruvate [ [14, 16, 17] ], which is incompatible with stable-isotope labeling and requires chemical pretreatment of the cell extract to stabilize the redox potential [ [14-16] ].


Systematic study of function across enzyme families has only recently been attempted and has not previously been applied to a representative sampling of an entire superfamily on this scale. A small, uncharacterized enzyme family, DUF849, was studied by Bastard et al. (3) using an integrated strategy for evaluating functional diversity. Based on screening of a set of representative enzymes (124 out of 900) against 17 putative substrates this study found that the family generally catalyzes the condensation of a β-keto acid with acetyl-CoA producing acetoacetate and CoA ester. Widespread catalytic promiscuity was described in the metallo-β-lactamase superfamily through the analyses of 24 enzymes against 10 catalytically distinct hydrolytic reactions where enzymes catalyzed on average 1.5 reactions in addition to their native activity (38). In an orthogonal study, screening of all 23 soluble HADSF members within a single species (E. coli) was performed with a set of 80 commercially available phosphorylated substrates (4). The results annotated the E. coli enzymes and demonstrated that HADSF members possess the capacity to hydrolyze a wide range of phosphorylated compounds, with 19 of the 29 enzymes tested showing a broad substrate range.

Our findings demonstrate, through a representative sampling of prokaryotic organisms in the HADSF, that the degree of substrate ambiguity is great and ubiquitous. This leads to the question, How might the presence of enzymes with broad substrate range contribute to the fitness of individual organisms and to complex microbial communities? Accumulation of metabolites may disrupt metabolic flux or metabolites themselves may prove toxic to organisms (39). Broad substrate overlap between HADSF members may provide a safeguard against such deleterious metabolic effects. Overlap in function may serve to provide sufficient turnover (without optimizing efficiency of any one enzyme) to metabolize large pools of similar substrates. Substrate overlap may also aid in the evolution of new metabolic function in response to environmental challenges. Promiscuity increases flexibility during evolution in response to changes in the substrate pool. During evolution, if there is a variation in the substrate pool, it is more facile for mutations to change the substrate specificity spectrum of broad, overlapping enzymes—that is, divergence before gene duplication (40, 41).

The finding that HADSF members with a minimal or no cap insertion (type C0) are more specific than those with domain insertions (types C1, C2A, or C2B, Fig. 4 B and C) is surprising because the cap domain provides the specificity determinants in the HADSF (10). The expectation might be that in C0 members there are few enzyme residues that interact with the substrate-leaving group and thus the leaving group could vary in size, shape, and electrostatic surface, producing an enzyme with broad specificity. However, an alternative model explains the observation made here. The substrate-leaving group structure may be limited by the necessity of acting as a barrier between bulk solvent and the active-site Mg 2+ cofactor, nucleophilic Asp, and Asp general acid/base, replacing the cap domain in this role. Indeed, structures of the C0 HADSF members Scp-1 (35) and GmhB (42) show that the fit between enzyme and substrate does not allow a probe the size of water to enter the active site (see SI Appendix, Fig. S22). Conversely, in enzymes with cap domains there is an enclosed active site and a number of enzyme residues from the cap specificity loop(s) are available to make interactions with different substrates, allowing evolution of activity against multiple substrates. Additionally, the mobility of the cap domain with respect to the core domain may allow access to different substrates of varying size.

The concept that domain insertions provide the raw material for the evolution of specificity determinants (including the residues that allow substrate ambiguity) is supported by recent findings. The directed evolution and bioinformatics analysis of a lactamase supports the model that enzymes are more capable of catalyzing multiple reactions when the active site is composed of loops juxtaposed to, but separate from, the core scaffold (43). Domain insertion can be considered an extreme example of this observation. The facilitation of substrate ambiguity via domain insertion can be a widespread phenomenon in protein evolution, because it has been estimated that ∼10% of insertion events are domain insertions (44). Overall, our findings are consistent with the concept that domain insertions act to drive the evolution of new functions. The widespread substrate ambiguity in the C1- and C2-type HADSF members observed herein may be a vestige of the evolutionary process. Previous work has shown that in the process of in vitro evolution selection pressure toward increasing one activity results in “generalists” that show multiple activities that were not selected for (45).

Materials and Methods

Plants, growth conditions and sampling

The work was carried out using Arabidopsis thaliana L. (Heynh) (ecotypes Ler, Col-O and Ws-2), the NASC <"type":"entrez-nucleotide","attrs":<"text":"N92274","term_id":"1264583","term_text":"N92274">> N92274 (pgi1𠄳), the pgi1𠄲 mutant [28], the aps1::T-DNA mutant (SALK_040155) [31], the pgm::T-DNA mutant (GABI_094D07), the gpt2::T-DNA mutant (GABI_454H06), pgi1𠄳 plants expressing either PGI1 or PGI1*, the pgi1𠄳/gpt2 and pgi1𠄲/gpt2 double mutants and the pgi1𠄲/sex1 and pgi1𠄳/sex1 double mutants. The pgi1𠄲/sex1 and pgi1𠄲/gpt2 double mutants were confirmed by PCR using the oligonucleotide primers listed in S1 Table. The 35S-PGI1 and 35S-PGI1* plasmid constructs utilized to produce PGI1 or PGI1* expressing pgi1𠄳 plants were produced as illustrated in S7 Fig. pgi1𠄳 plants expressing GBSS-GFP were produced using the 35S-GBSS-GFP plasmid construct [105] whereas pgi1𠄲 plants expressing GBSS-GFP were produced using the 35S-GBSS-GFP* plasmid construct produced as illustrated in S8 Fig. The plasmid constructs were transferred to Agrobacterium tumefaciens EHA105 cells by electroporation and utilized to transform Arabidopsis plants according to [106]. Transgenic plants were selected on the appropriate antibiotic-containing selection medium.

Unless otherwise indicated plants were cultured in soil in growth chambers under the indicated photoperiodic conditions (light intensity of 90 μmol photons sec -1 m -2 ) and at a constant temperature of 22ଌ. Harvested source leaves were immediately freeze-clamped and ground to a fine powder in liquid nitrogen with a pestle and mortar.

To analyze the effects of exogenously applied CKs on starch content plants were grown in vitro on MS agar plates at a constant temperature of 22ଌ under LD conditions. Three-weeks old plants were then transferred to MS agar plates containing the indicated concentrations of tZ. After two additional days leaves were harvested, and starch content was measured as described below.

Enzyme assays

One g of the frozen powder (see above) was resuspended at 4ଌ in 3 ml of 100 mM HEPES (pH 7.5), 2 mM EDTA and 2 mM dithiothreitol, 1 mM PMSF and 10 ml/L protease inhibitor cocktail (Sigma P9599), and centrifuged at 14,000 x g for 20 min. The supernatant was desalted by ultrafiltration on Vivaspin 500 centrifugal concentrator (Sartorius) and the protein extract thus obtained was assayed for enzymatic activities. AGP and UGP activities were measured following the two-step assay method described in [48]. PGI and SuSy were measured as described in [24] and [107], respectively. Adenylate kinase was assayed in the two directions of the reaction as described in [108] using an HPLC system (Waters corporation) fitted with a Partisil 10-SAX column. PGM and acid invertase were assayed as described in [29] and [109], respectively. Rubisco activity was measured according to [110]. Amylolytic activities were assayed as described in [111]. PPase and SPS were measured as described in [74]. SS activity was measured in two steps: (1) SS reaction and (2) measurement of ADP produced during the reaction. The SS assay mixture contained 50 mM HEPES (pH 7.5), 6 mM MgCl2, 3 mM dithiothreitol, 1 mM ADPG and 3% glycogen. After 5 min at 37ଌ reactions were stopped by boiling the assay mixture for 2 min. ADP was measured by HPLC on a Waters Associate’s system fitted with a Partisil-10-SAX column. One unit (U) is defined as the amount of enzyme that catalyzes the production of 1 μmol of product per min.

Non-reducing western blot analyses of AGP

For non-reducing western blots of AGP, 50 mg of the homogenized frozen material (see above) was extracted in cold 16% (w/v) TCA in diethyl ether, mixed, and stored at -20ଌ for at least 2 h as described in [48]. The pellet was collected by centrifugation at 10,000 x g for 5 min at 4ଌ, washed 3 times with ice-cold acetone, dried briefly under vacuum, and resuspended in 1x Laemmli sample buffer containing no reductant. Protein samples were separated on 10% SDS-PAGE, transferred to nitrocellulose filters, and immunodecorated by using antisera raised against maize AGP as primary antibody [48], and a goat anti-rabbit IgG alkaline phosphatase conjugate (Sigma) as secondary antibody.

Chromatographic separation of cytPGI and pPGI

Chromatographic separation of the two PGI isoforms was conducted using an AKTA FPLC from Amersham Pharmacia Biotech. Protein extracts of WT and pgi1𠄳 leaves (see above) were loaded onto a HiLoad 16/10 Q-sepharose HP anion exchange column (Amersham Pharmacia Biotech) equilibrated with 50 mM HEPES (pH 7.5). After washing the column, the adsorbed proteins were eluted with a linear 0𠄰.8 M NaCl gradient in 50 mM HEPES (pH 7.5). The flow rate was 5 ml/min and 2.5 ml fractions were collected. Fractions were analyzed for PGI activity as described above.

Native gel assay for PGI activity

PGI zymograms were performed as described in [29]. Protein extracts (see above) of both WT and pgi1 leaves were loaded onto a 7.5% (w/v) polyacrylamide gel. After electroforesis gels were stained by incubating in darkness at room temperature with 0.1 M Tris-HCl (pH 8.0), 5 mM F6P, 1 mM NAD + , 4 mM MgCl2, 0.2 mM methylthiazolyldiphenyl-tetrazolium bromide (Sigma M5655) and 0.25 mM phenazine methosulfate (Sigma P9625) and 1 U/mL of G6P dehydrogenase from Leuconostoc mesenteroides (Sigma G8404).

Analytical procedures

For determination of metabolites content, fully expanded source leaves of 30 days after germination (DAG) plants were harvested at the indicated illumination period, freeze-clamped and ground to a fine powder in liquid nitrogen with a pestle and mortar. ADPG content was measured by HPLC-MS/MS as described in [55]. For measurement of sucrose, glucose and fructose, a 0.1 g aliquot of the frozen powder was resuspended in 1 mL of 90% ethanol, left at 70ଌ for 90 min and centrifuged at 13,000 x g for 10 min. For measurement of G6P, F6P and G1P 0.5 g aliquot of the frozen powdered tissue was resuspended in 0.4 ml of 1 M HClO4, left at 4ଌ for 2 h and centrifuged at 10,000 x g for 5 min. The supernatant was neutralized with K2CO3 and centrifuged at 10,000 x g. Sucrose, glucose, fructose, F6P, G6P and G1P from supernatants were determined by HPLC with pulsed amperometric detection on a DX-500 Dionex system. NADP(H) and NAD(H) were measured as described in [112]. Starch was measured by using an amyloglucosydase�sed test kit (Boehringer Mannheim, Germany). For measurement of adenine nucleotides a 0.5 g aliquot of the frozen powder was resuspended in 0.4 ml HClO4, left at 4ଌ for 2 h and centrifuged at 10,000 x g for 5 min. The supernatant was neutralized with K2CO3 and centrifuged at 10,000 x g. Nucleotides content in the supernatant was measured by HPLC (Waters corporation) fitted with a Partisil 10-SAX column as described in [113]. Recovery experiments were carried out by the addition of known amounts of metabolites standards to the frozen tissue slurry immediately after addition of extraction solutions.

For determination of CKs levels, aliquots of the frozen leaves (see above) were lyophilized and CKs were quantified according to the method described in [114].

Iodine staining

Leaves harvested at the end of the light period were fixed by immersion into 3.7% formaldehyde in phosphate buffer. Leaf pigments were then removed in 96% ethanol. Re-hydrated samples were stained in iodine solution (KI 2% (w/v) I2 1% (w/v)) for 30 min, rinsed briefly in deionized water and photographed.

Gas exchange determinations

Fully expanded apical leaves were enclosed in a LI-COR 6400 gas exchange portable photosynthesis system (LI-COR, Lincoln, Nebraska, USA). The gas exchange determinations were conducted at 25ଌ with a photosynthetic photon flux density of 350 μmol m -2 s -1 . An was calculated using equations developed by [115]. g s values were determined as described in [116]. From the A/Ci curves, Vcmax, TPU and Jmax were calculated according to [117]. To avoid miscalculation of An and Ci due to leakage into the gasket of the gas analyzer, we performed CO2 response curves using an empty chamber. The values obtained for An and Ci in the empty chamber were compared with those of the chamber filled with a leaf and substracted from the values obtained with the empty chamber. ETR values were calculated according to [118] as ΦPSII x PPFD x 0.84 x 0.5, where PPDF is the photosynthetic photon flux density incident on the leaf, 0.5 was used as the fraction of excitation energy distributed to PSII [119] and 0.84 as the fractional light absorbance [120]. The rate of mitochondrial respiration in the dark was determined by measuring the rate of CO2 evolution in the dark.

Real-time quantitative PCR

Total RNA was extracted from leaves using the trizol method according to the manufacturer’s procedure (Invitrogen). RNA was treated with RNAase free DNAase (Takara). 1.5 μg RNA was reverse transcribed using polyT primers and the Expand Reverse Transcriptase kit (Roche) according to the manufacturer’s instructions. Real time quantitative PCR reaction was performed using a 7900HT sequence detector system (Applied Biosystems) with the SYBR Green PCR Master Mix (Applied Biosystems) according to the manufacturer’s protocol. Each reaction was performed in triplicate with 0.4 μL of the first strand cDNA in a total volume of 20 μL. The specificity of the PCR amplification was checked with a heat dissociation curve (from 60ଌ to 95ଌ). Comparative threshold values were normalized to 18S RNA internal control. The specificity of the obtained RT-PCR products was controlled on 1.8% agarose gels. Primers used for RT-PCRs of PGI1, BAM1, BAM2, BAM3 and BAM5 are listed in S2 Table.

Confocal microscopy

Subcellular localization of GFP-tagged GBSS was performed using D-Eclipse C1 confocal microscope (NIKON, Japan) equipped with standard Ar 488 laser excitation, BA515/30 filter for green emission and BA650LP filter for red emission.

Light and electron microscopy

Light microscopy and TEM analyses were carried out essentially as described in [105]. Briefly, small pieces (2 mm 2 ) of leaves were immediately fixed by submersion in a solution of 3% glutaraldehyde (v/v) in 0.05 M sodium cacodylate buffer, pH 7.4 (3 h at 4ଌ, under vacuum). After fixing, the specimens were washed in a cacodylate buffer (0.05 M sodium cacodylate, 1% sucrose), three times for 30 min each at 4ଌ, and post-fixed with a solution of 1% osmium tetroxide in the above cacodylate buffer (overnight, 4ଌ). After two washes, 30 min each, at 4ଌ with the same cacodylate buffer, the samples were dehydrated in an ethanol series and progressively embedded in LR White resin (London Resin Co., Reading, UK). Semithin (1 μm) sections were stained with 1% (w/v) toluidine blue in aqueous 1% sodium borate for direct observation with a Zeiss Axiophot photomicroscope (Zeiss, Oberkochen, Germany). Ultrathin (70� nm) sections for TEM were constructed with 2% aqueous uranyl acetate and lead citrate. Observations were performed with a STEM LEO 910 electron microscope (Oberkochen, Germany) at 80 kV, equipped with a Gatan Bioscan 792 camera (Gatan, Pleasanton, CA, USA).

The Bottom Line: Increased Research Productivity

When asked what having the NGC system in his lab will enable his group to do in the future, Fraser summarizes it thus: “Throughput and reliability are extremely important in a lab with many researchers using the same instrument. The NGC system will boost our capacity to purify multiple proteins more quickly. We used to do four runs a day, now we can do five — perhaps a 25% increase in throughput — because it is much quicker to switch between users.” This enhanced capacity promises to relieve a major bottleneck to research productivity in many labs, a sentiment echoed by senior proteomics researchers, who have often cited chronic problems with chromatography systems as a major source of downtime.

Although research at a basic level such as that conducted in the Fraser lab is typically far removed from technological applications, when asked to speculate on the scope of possible uses for the knowledge obtained in their work, Fraser replies that the study of translation mechanisms could have numerous therapeutic uses. For example, “Understanding how viruses hijack ribosomes to translate their mRNA may lead to the development of drugs that can specifically inhibit viral translation (Fraser and Doudna, 2007). Our studies could also help elucidate the role of translational regulation in cancer.” In a world living with an HIV epidemic with no cure in sight and facing increasing cancer mortality, such technologies could be of great benefit to people all across the globe.

Example 3

Objective: Identification of the HMGB1 family in the skin extract and examination of bone marrow mesenchymal stem cell-inducing activity

Method: Whether or not the neonatal mouse skin extract contained the HMGB protein family was confirmed using the Western blot method. Ten μl of the skin extract obtained in [Example 2] was used as a sample and subjected to SDS-PAGE electrophoresis. The proteins separated within the gel were transferred onto a PVDF membrane using a blotting device (ATTO). The membrane was incubated with PBS containing 3% skim milk and 0.1% Tween 20 (S-T-PBS) at room temperature for 1 hour, and then was allowed to react with each of rabbit anti-mouse HMGB1 antibody, rabbit anti-mouse HMGB2 antibody, or rabbit anti-mouse HMGB3 antibody which were diluted 1000-fold with S-T-PBS, at 4° C. for 16 hours. After the reaction, the PVDF membrane was washed with S-T-PBS five times for 5 minutes. Then, the PVDF membrane was incubated with 2000-fold diluted (diluted with S-T-PBS) peroxidase labeled goat anti-rabbit IgG antibody (GE Healthcare) at 25° C. for 1 hour. Further, after washing with S-T-PBS five times for 5 minute, the PVDF membrane was allowed to react with ECL Western Blotting Detection System (GE Healthcare). The ECL film was exposed and developed to detect the presence of HMGB1, HMGB2, and HMGB3 proteins.

RNA was extracted from the skin of neonatal mouse using Trizol (invitrogen), and further cDNA was synthesized using SuperScript III cDNA synthesis kit (Invitrogen). Using this cDNA as a template, cDNAs of HMGB1, HMGB2, and HMGB3 were amplified using the PCR (polymerase chain reaction) method. The cDNAs were inserted into the plasmid vector pCAGGS for expressing proteins in mammalian cells, such that proteins with an additional Flag tag sequence (Asp-Tyr-Lys-Asp-Asp-Asp-Lys SEQ ID: 18) at the N terminus of the amino acid sequence could be expressed. These plasmid vectors were introduced into HEK293 (Human embryonic kidney derived culture cell line) and cultured for 48 hours to express the proteins. Cells expressing each of the HMGB1, HMGB2, and HMGB3 proteins and the culture supernatant were incubated at 4° C. for 16 hours, which was then centrifuged at 4400 g for 5 minutes to collect the supernatant. 100 μL of the anti-Flag antibody gel (Sigma) was mixed into 50 mL of this supernatant, and was then incubated at 4° C. for 16 hours. Centrifugation was then performed to collect the gel, and washed with PBS five times. Further, the protein was eluted using 3× Flag peptide (final 100 μg/ml). Expressions of recombinant proteins were observed by the Western blot method using 1000-fold diluted (diluted with S-T-PBS) mouse anti-Flag antibody and 2000-fold diluted (diluted with S-T-PBS) peroxidase-labeled anti-mouse IgG antibody (GE Healthcare). The mouse bone marrow mesenchymal stem cell migration activity in these purified recombinant proteins was assessed in the same manner as in [Example 2] using a Boyden chamber. Moreover, in order to observe the in vivo drug efficacy of the HMGB family, the dorsal skin of 8-week-old C57BL/6 mice was cut out in a circle having a diameter of 8 μm to prepare cutaneous ulcer models. Purified HMGB1, HMGB2, and HMGB3 (100 ng) were each mixed with the same amount of hyaluronic acid solution having a concentration of 1 g/100 mL of PBS, and 100 μL of it was administered to the ulcer surface. The ulcer surface was covered with a transparent adhesive wound dressing/protective material Tegaderm (3M Healthcare) to avoid drying, and the wound area was measured over time to determine the therapeutic effect.

Further, to examine whether or not the human skin extract and the purified human HMGB1 has an activity to allow migration of human bone marrow mesenchymal stem cells, a Boyden chamber was used in the same manner as in [Example 2] for assessment. A human skin having an area of 1 cm 2 was immersed in 1 ml PBS, and then was incubated at 4° C. for 16 hours and subsequently centrifuged at 440 at 4° C. for 10 minutes. The supernatant alone was collected to be used as a human skin extract. Moreover, human bone marrow mesenchymal stem cells (Cambrex) were used as the cells to be placed in the upper chamber of the Boyden chamber (as a result of surface antigen analysis by flow cytometry, these cells have been confirmed to be CD105-positive, CD166-positive, CD29-positive, CD44-positive, CD34-negative, and CD45-negative. They have also been found to differentiate into adipocytes, chondrocytes, and bone cells by differentiation induction tests). Moreover, 100 ng/well of human HMGB1 (R&D) and human skin extract diluted 10-fold with PBS and were placed in the lower chamber. PBS was used as a control.

Result: As a result of Western blotting, bands of HMGB2 and HMGB3 were detected as well as the HMGB1 band. Therefore, the neonatal mouse skin extract was confirmed to contain the family proteins, HMGB2 and HMGB3, besides HMGB1 (FIG. 15). Expression vectors of HMGB1/HMGB2/HMGB3 having a Flag tag added at the N-terminus of each protein, were prepared (FIG. 16). These expression vectors were introduced into HEK293 cells, and the expressed proteins were purified using the Flag tag, and Western blotting was carried out to observe these proteins (FIG. 17). The mouse bone marrow mesenchymal stem cell migration activity was measured using these purified proteins, and the activity was confirmed in all of the proteins (FIG. 18). The ulcer area produced in the back of the mouse was measured every 7 days, and a significant effect on reducing ulcer area was confirmed in the HMGB1, 2, and 3 treatment group, as compared to the non-treatment group (FIG. 19). Similar to the mouse case, human HMGB1 and the human skin extract were revealed to have human bone marrow mesenchymal stem cell migration activity (FIG. 20).

Discussion: HMGB2 and HMGB3 are known as proteins having high homologies to HMGB1. These proteins are also expected to have properties similar to HMGB1. It was confirmed that HMGB2 and HMGB3 of the HMGB1 family are also produced from the extract of the free skin section. Further, HMGB1/HMGB2/HMGB3 recombinant proteins were produced, and their in vitro bone marrow mesenchymal stem cell migration activity and the in vivo therapeutic effect on a cutaneous ulcer were also confirmed. It was revealed that the HMGB family (HMGB1/HMGB2/HMGB3) and the recombinant HMGB family in the neonatal mouse free skin section have a bone marrow mesenchymal stem cell-inducing activity and an activity of locally inducing bone marrow-derived stem cells which are differentiatable into epithelium, and that the thus induced bone marrow-derived cell group differentiates into various cells such as epidermal keratinocytes, hair follicles, and fibroblasts in the damaged tissue to promote the recovery of the damaged tissue. Moreover, since bone marrow mesenchymal stem cells are multipotent stem cells, the present inventors believe that therapeutic effects can also be expected in the same manner by systematic administration or local administration of the HMGB family to treat damaged states in other tissues, for example, tissue damages such as brain injury, myocardial infarction, and bone fracture.

Moreover, it is known that, between human and mouse, amino acid sequence homology for HMGB1 is 98% (213/215), 96% (202/210) for HMGB2, and 97% (195/200) for HMGB3. Therefore, human HMGB and mouse HMGB are considered to have similar activities, and the results of the present Examples revealed that human skin extract and human HMGB1 have bone marrow mesenchymal stem cell-inducing activities in a manner same as those of mouse skin extract and mouse HMGB1.


A.V., S.M., T.M., L.V., C.L. were supported by CNRS (Mission pour l’interdisciplinarité, Agromics 2014–2016) this work was submitted to fulfill the requirements for a doctorate of biology at ED341-E2M2 from Université de Lyon, granted from the French Ministère de l’Education Nationale, de l’Enseignement Supérieur et de la Recherche (to T.M.) this work has benefited from the I2BC crystallization and protein–protein interactions platforms supported by FRISBI [ANR-10-INSB-05-01] Cell and Tissue Imaging (PICT IBiSA), Institut Curie, member of the French National Research Infrastructure France-BioImaging [ANR10-INBS-04]. Funding for open access charge: CNRS.

Conflict of interest statement. None declared.

An Economical and Versatile High-Throughput Protein Purification System Using a Multi-Column Plate Adapter

A multi-column plate adapter allows chromatography columns to be interfaced with multi-well collection plates for parallel affinity or ion exchange purification providing an economical high throughput protein purification method. It can be used under gravity or vacuum yielding milligram quantities of protein via affordable instrumentation.

An economical and versatile high throughput protein purification system using a multi column plate adapter. Hi, protein purification is imperative to the study of protein structure and function. And it's usually used in combination with biophysical techniques.

This is especially important in the era of functional proteomics that require a high-throughput protein production. We hypothesized that a multi-column plate adapter, the MCPA, can interface multiple chromatography columns of different resins with multi-well plates for parallel purification. With this adapter, we have developed an economical and versatile method of protein purification that can be used under gravity or vacuum rival in the speed of an automated system.

Here we show this method for nickel affinity chromatography and ion exchange chromatography. Assemble the MCPA by placing a punctured sealing mat onto a long drip plate, then insert the desired number of columns into the holes of the sealing mat. Place an open collection plates in the base of the manifold and close with the top of the manifold.

Attach tubing and place the assembled MCPA with columns on the top. Pipet 1.2 mil of the nickel-NTA solution into the columns. Whilst pipetting, ensure that the beads remain fully mixed.

Switch on the pump and run the 20%ethanol through the columns and into the collection plate below. Turn off the pump once the liquid is run through. Dispose of the contents of the collection plate into a waste box.

Add three resin volumes of EDTA buffer to all the columns. Turn on the pump and run the liquid through. Now wash the columns with three resin volumes of 0.5 molar sodium hydroxide buffer.

Wash the columns with four resin volumes of 100 millimolar nickel sulfate, then 10 resin volumes of Mili-Q water. If a column is running slower, push the liquid through with the syringe plunger. Pour out the contents of the collection plates into a waste box.

Then wash the columns twice with four resin volumes of 10 millimolar imidazole wash buffer and empty the collection plates. If multiple samples are being purified, replace the open collection plates with a 48-well plates. With the vacuum off, load the lysates into the columns.

Use a thin plastic stirrer to gently mix the beads and the lysates in the column to maximize binding. Turn on the pump and run the liquid through. If a column becomes blocked, transfer the lysate resin mixture to a column with a fresh filter.

Freeze the collection plates to contain your flow-through. Replaced with an open collection plate and then wash the columns with nine resin volumes of 10 millimolar imidazole wash buffer. Repeat this step four or five times.

To avoid overflow, periodically empty the waste plates. Replace the open collection plates with a 96-well plate, ensuring A1 is in the top left corner. Pipet one resin volume of denaturing elution buffer into the columns.

Run the liquid through and check the collection plate to ensure that there is no contamination between the wells. For the next dilution, repeat the previous steps and collect in a fresh collection plate. Take a 50 microliter aliquot of each elution for the purity and concentration analysis.

Next, wash the columns with two milliliters of 20%ethanol. Add another two mil of 20%ethanol to the columns and use the fresh thin plastic stirrer to mix up the beads in the ethanol before transferring to a 50 mil tube for storage at four degrees Celsius. Assemble the MCPA in vacuum manifold with an open collection plate and syringe plungers in all of the columns.

Remove all of the syringe plungers from the front row. Remove the rubber gasket from a five mil syringe plunger and then pierce a hole in the center. Then push the bottom of an open column through the punctured rubber gasket.

Repeat these steps and insert into the front row of columns. Ensure that the Q sepharose beads are fully mixed. And with the blue pipette tip that is cut two centimeters from the bottom, pipet 800 microliters of the Q sepharose beads into the open columns.

Once the beads have settled to the bottom of the columns, switch on the vacuum pump enough to run the 20%ethanol through. Wash the columns twice with two mil of 10 millimolar Tris. Turn off the pump just before the Tris buffer has run through to prevent the resin drying out.

Runoff can be discarded and replace with a 48-well collection plates. Transfer all samples to be purified to the Q Sepharose columns in the first row and use a thin plastic stirrer to mix the samples and the beads for around two minutes before turning on the vacuum pump. Store or freeze your flow-through for later analysis.

With an open collection plate, wash the columns twice with two mil of 10 millimolar Tris. Replace the open collection plates with the 96-well plate. The first elution fraction can be collected in the first row or in the second row.

To elute in the second row, remove the syringe plungers from these positions. Move the Q Sepharose columns into these positions and then place syringe plungers into the open columns in the first row. Pipet one milliliter of the first salt concentrations into the Q Sepharose columns.

Turn on the pump and collect the elution. To collect the next elution, remove the syringe plungers from the next positions, move the Q Sepharose columns into these positions and then replace the plungers in the previous positions. Repeat these steps for each successive salt concentration used to elute.

Ensure that a new collection plate is used for every four elutions and that every plate is labeled and stored. With an open collection plates, wash the columns with two mil of four molar sodium chloride, 10 millimolar Tris. Pipet two mil of 20%ethanol and run this through until it is just above the beads and then seal the columns for storage.

As an example, the MCPA has successfully purified 14 AbpSH3 mutants in denaturing conditions by a nickel-NTA. A small contaminant can be seen at 25 kilodaltons, though the protein is still largely pure. The small contaminants seen in denaturing conditions is removed in native conditions.

This was shown when 11 different SH3 domains were purified. This shows that the MCPA can be used for comparison of purification conditions. AbpSH3 can be separated from the majority of contaminants via ion exchange purification from lysates.

Good yields of considerably pure AbpSH3 protein were recovered with various higher molecular weights contaminants. Purification of samples post-nickel-NTA using ion exchange with the MCPA has successfully removed these high-weight contaminants. Though there is still a slight contamination, the fractions are still largely pure and have yielded good biophysical data using NMR and thermal chemical denaturation assays.

In conclusion, our results show that our hypothesis is correct and that the MCPA can successfully purify milligram quantities of proteins using different chromatography techniques. The MCPA system can be configured in multiple ways within the same runs to optimize purification conditions. The setup is simple, cheap, and easy to train inexperienced users, especially in labs that do not routinely purify proteins.


Figure 1. Polymer p(SS-co-PEGMA) stabilizes FGF2 as a conjugate and pVS facilitates FGF2-receptor binding when added as an excipient. When combined into a block copolymer, p(SS-co-PEGMA)-b-VS, the new conjugate both stabilizes FGF2 and increases protein activity. Protein structure modified from PDB 1CVS using PyMOL software.

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