PEDS Advance Access originally published online on February 20, 2008
Protein Engineering Design and Selection 2008 21(5):303-310; doi:10.1093/protein/gzn005
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Thermostability promotes the cooperative function of split adenylate kinases
Department of Biochemistry and Cell Biology, Rice University, Houston, TX 77251, USA
1 To whom correspondence should be addressed. E-mail: joff{at}rice.edu
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Abstract |
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Proteins can often be cleaved to create inactive polypeptides that associate into functional complexes through non-covalent interactions, but little is known about what influences the cooperative function of the ensuing protein fragments. Here, we examine whether protein thermostability affects protein fragment complementation by characterizing the function of split adenylate kinases from the mesophile Bacillus subtilis (AKBs) and the hyperthermophile Thermotoga neapolitana (AKTn). Complementation studies revealed that the split AKTn supported the growth of Escherichia coli with a temperature-sensitive AK, but not the fragmented AKBs. However, weak complementation occurred when the AKBs fragments were fused to polypeptides that strongly associate, and this was enhanced by a Q16L mutation that thermostabilizes the full-length protein. To examine how the split AK homologs differ in structure and function, their catalytic activity, zinc content, and circular dichroism spectra were characterized. The reconstituted AKTn had higher levels of zinc, greater secondary structure, and >103-fold more activity than the AKBs pair, albeit 17-fold less active than full-length AKTn. These findings provide evidence that the design of protein fragments that cooperatively function can be improved by choosing proteins with the greatest thermostability for bisection, and they suggest that this arises because hyperthermophilic protein fragments exhibit greater residual structure compared to their mesophilic counterparts.
Keywords: adenylate kinase/hyperthermophile/protein design/protein fragment complementation/protein thermostability
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Introduction |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionFundingAcknowledgementsReferences |
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In nature, protein sequence diversity is created through a variety of processes, including gene fission. The frequency of fission has been quantified in several organisms (Snel et al., 2000
; Pasek et al., 2006), but its effects on protein function cannot yet be predicted a priori, limiting our ability to design split proteins with tailored functions for applications in systems (Michnick, 2003) and synthetic (Giesecke et al., 2006) biology. Single-domain proteins having diverse topologies have been bisected into polypeptide fragments that cooperatively function, including adenylate kinase (Monnot et al., 1987, Saint Girons et al., 1987), aminoacyl-tRNA synthetase (Shiba and Schimmel, 1992), chymotrypsin inhibitor-2 (de Prat Gay et al., 1994), dihydrofolate reductase (Gegg et al., 1997), green fluorescent protein (Cabantous et al., 2005), imidazoleglycerol phosphate synthase (Hocker et al., 2001), and ribonuclease S (Kato and Anfinsen, 1969). These split proteins have been used to test questions about protein evolution, folding, solubility and structure–function relationships. In addition, studies with non-functional split proteins have revealed that fragment complementation can often be enhanced by fusing the inactive polypeptides to proteins that associate and promote oligomerization (Johnsson and Varshavsky, 1994; Pelletier et al., 1998; Remy and Michnick, 1999; Magliery et al., 2005; Nyfeler et al., 2005; Wilson et al., 2004; Remy and Michnick, 2006). Such assisted protein fragment complementation has been useful for probing protein–protein interactions in cellular compartments (Nyfeler et al., 2005, Spotts et al., 2002) and organisms (Paulmurugan et al., 2002; Singh et al., 2006) inaccessible by the classical yeast-two hybrid assay (Fields and Song, 1989).
Combinatorial protein engineering studies have shown that for a given enzyme, a diverse array of cut points can yield polypeptides that associate and form functional heterodimers (Ostermeier et al., 1999). Non-disruptive breaks have been found within all types of structural elements (i.e., in loops, 
-helices, and β-sheets) and between residues that constitute active sites (Ostermeier et al., 1999). What is poorly understood is whether bisection of protein orthologs at the same site leads to polypeptide pairs with similar levels of parent-like structure and function. Directed evolution studies examining the effects of other mutational processes on the retention of protein function suggest that protein thermostability will influence the activity of split proteins. Among homologous proteins, those with the highest thermostability produce the largest fraction of functional variants upon random mutation (Bloom et al., 2005; Bloom et al., 2006; Besenmatter et al., 2007). Furthermore, the fraction of functional variants in chimeric libraries created by recombination can be enhanced by incorporating mutations that are known to stabilize the structure of the proteins recombined (Meyer et al., 2006).
To establish whether protein thermostability affects the cooperative function of fragments that arise when a polypeptide backbone undergoes fission, we evaluated the structure and function of protein fragments derived from protein orthologs having different thermostabilities. As our model system, we chose adenylate kinases, ubiquitous phosphotransferases that maintain cellular adenine energy charge by catalyzing the reversible reaction of ATP + AMP
2ADP.
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Materials
Escherichia coli (E. coli) XL1-Blue, and Rosetta 2 cells were from Stratagene and EMD Biosciences, respectively. Enzymes for DNA manipulation were obtained from Roche Biochemical, New England Biolabs, and Promega. Synthetic oligonucleotides were obtained from Fischer Scientific, and pET vectors were from EMD Biosciences. Bacterial growth media components were from BD Biosciences, and all other reagents were from Sigma-Aldrich.
The gene encoding AKBs was amplified from genomic DNA using PCR with Vent DNA polymerase and cloned into pET21d using NcoI and HindIII restriction enzymes to create pBsAK. Previously described pET vectors for expressing AKTn (pTNAK2::Km) and AKBs with Q16L and Q199R mutations (designated pBsAKQ16L/Q199R herein) were used for studies involving full-length proteins (Vieille et al., 2003, Counago et al., 2006).
Vectors for producing AKBs fragments with associating polypeptide tags fused to their N-terminus were generated by chemically synthesizing genes encoding the IAAL-E3 and IAAL-K3 peptides (Litowski and Hodges, 2002) followed by a flexible linker of sequence GASGGGSSGGHM and ligating these to PCR amplified gene fragments encoding AKBs residues 1–76 (AK-N) and 77–217 (AK-C), respectively. In each of these gene fragments, three unique restriction sites (NheI, XhoI, and NdeI) were incorporated within codons of the linker. The IAAL-E3–AK-N and IAAL-K3–AK-C gene fusions were cloned into pET21d and pET24d using NcoI and HindIII restriction enzymes to generate pB-E3-N and pB-K3C, respectively.
Vectors for producing AKBs fragments alone (pB-N and pB-C) were created by converting the NcoI restriction sites in pB-E3N and pB-K3C to NdeI using QuikChange mutagenesis, removing the gene fragments encoding the IAAL-E3 and IAAL-K3 peptides and linker by NdeI digestion, and circularizing the vector by ligation. Q16L and Q199R mutations were introduced into pB-N, pB-E3N, pB-C, and pB-K3C by Quikchange mutagenesis to create pB-NQ16L, pB-E3NQ16L, pB-CQ199R, and pB-K3CQ199R, respectively.
AKTn gene fragments corresponding to residues 1–79 (AK-N) and 80–220 (AK-C) were amplified from pTNAK2::Km using Vent DNA polymerase, and cloned in place of the genes encoding structurally-related Bacillus subtilis (B. subtilis) fragments in pB-N, and pB-C using NdeI and HindIII to create pTn-N and pTn-C, respectively. AKTn gene fragments were also cloned in place of AKBs fragments in pB-E3N and pB-K3C using NcoI and HindIII to create pT-E3N and pT-K3C, respectively. Vectors for expressing B. subtilis (pB-His-N and pB-His-C) and Thermotoga neapolitana (T. neapolitana) (pT-His-N and pT-His-C) AK fragments with N-terminal (His)6 tags were generated by subcloning AK gene fragments from pET21d (pB-N and pT-N) and pET24d (pB-C and pT-C) derived vectors into pET28b using NdeI and HindIII.
E. coli CV2 was used for all growth selections (Haase et al., 1989). Electrocompetent cells (designated CV2T7) were generated that harbor pTara, a vector that produces T7 RNA polymerase under control of the araBAD promoter element (Wycuff and Matthews, 2000). This vector constitutively produces T7 RNA polymerase when cells are grown on LB medium, thereby driving low level expression of transcripts controlled by T7 promoters. CV2T7 exhibited a reversion frequency of <1 in 107 cfu.
To assess complementation, CV2T7 were cotransformed by electroporation with plasmids encoding each AK fragment pair, and the transformed cells were grown at 30°C for 48 h on LB-agar plates containing 17 µg/ml chloramphenicol, 50 µg/ml ampicillin, and 50 µg/ml kanamycin. Studies with full-length AK homologs were performed similarly with only those antibiotics required to select for maintenance of pTara and the AK-containing plasmid. Colonies obtained from initial plates were streaked onto LB-agar plates lacking antibiotic and incubated at 40°C to assess AK complementation.
Colonies obtained from growth at 30°C were also used to inoculate 500 µl LB cultures in 96-well plates containing 17 µg/ml chloramphenicol, 50 µg/ml ampicillin, and 50 µg/ml kanamycin. These plates were grown for 24 h at 30°C while shaking at 250 rpm. A 96-pin replicator was used to transfer 1 µl of cells from each well into plates containing fresh LB medium. These plates were grown at 40°C in a humidified incubator shaking at 250 rpm, and the cellular density was evaluated by measuring absorbance at 600 nm.
Protein expression and purification
Rosetta 2 E. coli transformed with pB-His-N, pB-His-C, pT-His-N and pT-His-C were grown in LB at 37°C, induced with 1 mM IPTG at A600
1 and grown for 6 to 8 h at 25°C to allow for protein expression. Cells were harvested by centrifugation, resuspended in TND buffer (50 mM Tris pH 8.0, 300 mM NaCl, and 1 mM DTT) containing 25 mM imidazole, 0.1 mM PMSF, 1 mM MgCl2, 300 µg/ml lysozyme, 2 U/ml DNase, and a complete protease inhibitor cocktail tablet (Roche Applied Science). Resuspended cells were frozen at –80°C for 
2 h, thawed, and centrifuged at 40 k x g to remove cell debris. The cleared lysate was bound to NTA resin and washed with TND buffer containing 25 mM imidazole, 5 mM MgATP, and 12.5 µg/ml of denatured E. coli lysate to remove any ATP-dependent chaperones bound to the AK fragments (Rial and Ceccarelli, 2002). The denatured lysate was prepared from Rosetta 2 cells by boiling cleared cell lysate at 100°C for 10 min and then centrifugation at 14 k rpm to remove all aggregates. AK fragments were eluted using a linear gradient of 0 to 250 mM imidazole in TND buffer and further purified using a combination of ion-exchange and gel filtration chromatography. The purified proteins were concentrated in TND buffer and stored at –70°C. SDS–PAGE analysis of each purified protein revealed that they were all 95% homogeneous.
Rosetta 2 E. coli transformed with pTNAK2:Km was grown in LB at 37°C, induced with 0.1 mM IPTG at A6001 and grown for 18 h at 37°C to allow expression. Full length AKTn was purified using a combination of ion exchange, reverse phase, and size exclusion chromatography. AKTn was concentrated, dialyzed against TED buffer (50 mM Tris pH 8.0, 0.5 mM EDTA, and 1 mM DTT) and stored at –70°C.
The relative zinc content of AK fragments was determined by incubating 3 to 6 µM of each fragment with 100 µM PAR in 50 mM HEPES pH 7.5 and evaluating the release of zinc upon addition of 200 µM MMTS. The reaction was incubated at room temperature for 60 min, centrifuged to remove any precipitated protein, and changes in absorbance at 500 nm were monitored. The concentration of zinc was calculated using an extinction coefficient for the PAR–Zn complex (
500 = 60,423 M–1cm–1) that was obtained in HEPES pH 7.5 using ZnCl2 as a standard. The data reported represents the average of three replicates, each corrected for zinc levels in the buffer, with error corresponding to one standard error.
AK activity was determined using an endpoint assay that measures the amount of ADP formed in a reaction containing ATP and AMP (Vieille et al., 2003). Reactions (350 µl) containing HKM buffer (50 mM HEPES pH 7.4, 250 mM KCl, and 4 mM MgCl2) and 0.0005 to 1 µM protein were incubated at 40°C for 5 min prior to initiating the reaction by adding AMP and ATP to final concentrations of 1 mM. After 2 to 5 min, reactions were quenched by placing them on ice and adding P1,P5-di(adenosine-5) pentaphosphate (55 µl) to a final concentration of 0.25 mM. A mixture (150 µl) containing 1.85 mM phosphoenolpyruvate, 0.925 mM NADH, and 2.5 units of lactate dehydrogenase in HKM buffer was added to the quenched reaction, and NADH consumption was monitored by following increases in absorbance at 340 nm after the addition of 2.5 units of pyruvate kinase. The amount of NADH consumed was calculated using an 340 of 6,220 M–1cm–1, and the concentration of ATP consumed in the original reaction mixture was calculated as 0.793·[NADH]consumed.
Protein concentrations were assessed using the Bradford method with BSA as a standard. UV–Vis absorbance was measured using SpectraMax M2 microplate and Cary 50 spectrophotometers. Far-UV circular dichroism (CD) spectra were recorded with a Jasco J-810 spectrapolarimeter using a pathlength of 0.1 cm. All CD spectra were acquired with proteins that were dialyzed into TD buffer (20 mM Tris pH 8.0 and 0.5 mM DTT).
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Results |
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AK protein fragment design
To test whether thermostability affects protein fragment complementation, we have split AK orthologs from B. subtilis and T. neapolitana that exhibit a range of thermostabilities and compared their structure and function at 40°C. AKBs and AKTn are monomeric zinc-binding enzymes that are functional at 30°C (Vieille et al., 2003, Bae and Phillips, 2004). However, they differ in the temperature profiles for their activity. AKBs exhibits maximal activity at 
45°C and is largely inactive at 60°C (Bae and Phillips, 2004), whereas AKTn is maximally active at 80°C and retains almost 50% of its activity at 100°C (Vieille et al., 2003). An AKBs mutant (Q16L/Q199R) with intermediate thermostability was also used in these studies. This protein is maximally active at 60°C and exhibits a midpoint for thermal denaturation that is 14°C greater than AKBs (Counago et al., 2006) and 30°C less than AKTn (Vieille et al., 2003).
The AKBs structure shown in Fig. 1 illustrates the site of bisection and the polypeptide fragments used for complementation studies. All AK orthologs have been split to produce N-terminal fragments (designated AK-N) that contain the AMP binding site and C-terminal fragments (designated AK-C) that contain all of the residues involved in zinc coordination and the mobile LID that mediates substrate entry into the active site (Schulz et al., 1990; Muller et al., 1996). Both fragments contain residues from the hydrophobic core of the protein. This fragmentation site was chosen based on a previous study that showed that E. coli AKEc could be similarly cleaved at this site in vitro to create polypeptides that recover partial catalytic activity when reconstituted at 23°C (Saint Girons et al., 1987). However, AKEc was avoided herein because functional complementation was assayed in an E. coli strain with a temperature-sensitive AK (Haase et al., 1989). Complementation assays using non-native AK homologs are less likely than AKEc to lead to chromosomal recombination and reversion of the temperature-sensitive phenotype.
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AK fragment complementation
To initially compare the activities of split AK homologs, we examined whether they could complement the growth of E. coli CV2, a strain with a temperature-sensitive AK (Pro87Ser) that cannot support growth at 40°C (Haase et al., 1989). Genes encoding each AK-N and AK-C fragment were cloned into pET vectors with different antibiotic selection markers. In pET vectors, AK fragment expression is controlled by the T7 promoter, and all studies involving E. coli CV2 used competent cells that harbor pTara (designated CV2T7), a plasmid that constitutively produces T7 RNA polymerase when cells are grown on LB medium (Wycuff and Matthews, 2000).
To evaluate the relative activities of split AK homologs, CV2T7 were transformed with vectors for expressing three full-length AK homologs (pBsAK, pBsAKQ16L/Q199R, and pTNAK2) and their corresponding fragment pairs (pB-N/pB-C, pB-NQ16L/pB-CQ199R, and pT-N/pT-C). Cells were selected for growth at 30°C on LB-agar plates containing antibiotics that maintained the desired sets of plasmids, colonies obtained from selections were streaked on LB-agar plates lacking antibiotics, and CV2T7 growth was evaluated under selective (40°C) and non-selective (30°C) conditions.
Figure 2 shows the complementation of CV2T7 by full-length and fragmented AK homologs. Complementation occurred at 40°C after 24 h with CV2T7 harboring plasmids for producing T. neapolitana AK-N and AK-C fragments, similar to that obtained with CV2T7 expressing full-length AKTn, AKBs, and AKBsQ16L/Q199R. In addition, cells harboring the AKBsQ16L/Q199R fragment pair derived from the AK with intermediate thermostability appeared at a low density after 72 h. However, no growth occurred after 72 h with CV2T7 alone or CV2T7 expressing the split AKBs, although these cells were viable at 30°C.
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Assisted protein fragment complementation
Fragment complementation can frequently be improved by fusing the inactive fragments to proteins that associate (Johnsson and Varshavsky, 1994; Pelletier et al., 1998; Remy and Michnick, 1999; Wilson et al., 2004; Magliery et al., 2005; Nyfeler et al., 2005; Remy and Michnick, 2006). To test whether this strategy can promote AK-fragment function, we fused the AKBs fragments to the IAAL-E3 and IAAL-K3 peptides that strongly associate (KD = 70 nm) to form a heterodimeric coiled-coil (Litowski and Hodges, 2002) and evaluated whether these tagged AK fragments exhibit improved CV2T7 complementation. The pET vectors created for expressing these polypeptide fusions were designed so that each AK fragment has the associating polypeptides fused to their N-terminus through a twelve-residue linker that is predicted to be flexible (Linding et al., 2003).
Figure 3 shows the effect of IAAL-E3 and IAAL-K3 on the function of split AKBs. When fused to these associating tags, AK-NQ16L and AK-CQ199R complemented CV2T7 growth at 40°C after 24 h, as did AK-NQ16L and AK-C. In contrast, CV2T7 expressing the native AK-N and AK-C fusions exhibited weak complementation after 48 h, although these cells grew readily after 24 h at 30°C under non-selective conditions. A similar weak growth phenotype was observed at 40°C for CV2T7 harboring plasmids for producing AK-N and AK-CQ199R fused to associating tags.
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Complementation by hybrid fragments
To examine which AK fragments cooperatively function in liquid cultures, we evaluated CV2T7 complementation in 96-well microtiter plates for all possible AK-N and AK-C combinations. For these assays, colonies selected to grow at 30°C on LB-agar plates were used to inoculate 200 µl LB cultures in microtiter plates containing ampicillin, chloramphenicol, and kanamycin. After 24 h of growth at 30°C, these cultures were replicated into fresh microtiter plates lacking antibiotic and grown for 72 h in a humidified shaking incubator at 40°C. Complementation was evaluated at various times by measuring optical density at 600 nm.
Figure 4A shows the growth of CV2T7 transformed with all possible combinations of AK-N and AK-C fragments. As with the selections on LB-agar medium, complementation was observed when cells contained vectors for expressing T. neapolitana AK-N and AK-C. However, B. subtilis AK fragments containing the Q16L and Q199R mutations did not weakly complement growth in liquid medium as was observed on LB-agar plates. This indicates that the liquid growth selections are more stringent than those on agar plates.
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Figure 4B shows the results from complementation studies that used AK fragments fused to the IAAL-E3 and IAAL-K3 associating polypeptides. Again, the AKTn fragments complemented CV2T7, with maximal growth occurring within 24 h. In addition, complementation was observed for cells coexpressing B. subtilis AK-NQ16L and AK-C, B. subtilis AK-NQ16L and AK-CQ199R, and B. subtilis AK-NQ16L and T. neapolitana AK-C. These findings show that only AKBs fragments with the Q16L mutation can form a functional split protein when selection are performed in liquid medium, and they demonstrate that only AKBs fragments with this mutation can cooperatively function with an AKTn fragment.
To obtain AK fragments for in vitro studies, we have created IPTG-controlled expression plasmids that produce AK-N (pB-His-N and pT-His-N) and AK-C (pB-His-C and pT-His-C) fragments with a (His)6 affinity tag at their N-terminus. The addition of a His tag to AKTn fragments did not affect their ability to complement CV2T7 E. coli. Cells (108 cfu) cotransformed with pT-His-N and pT-His-C yielded a large number of colonies (105 cfu) at 40°C after 48 h.
AK-N and AK-C fragments were overexpressed in E. coli growing in LB medium and purified using a similar combination of NTA-affinity, anion-exchange, and size-exclusion chromatography in the absence of zinc chelators like EDTA. All fragments were soluble. Whereas the yields for the purified AK-C fragments were 
1.5 mg/l culture medium, the yields for AK-N fragments were 5-fold lower (0.3 mg/l).
To examine how fragmentation affects AK function, we have compared the activities of the split and full-length AK homologs in catalyzing the formation of ADP from ATP and AMP. Figure 5 shows the rates obtained at 40°C in the presence of AMP and ATP concentrations (1 mM) that that are >10-fold higher than the Km values reported for the full-length proteins (Glaser et al., 1992; Vieille et al., 2003). At this temperature, full-length AKTn and AKBs displayed similar rates of 1.5 x 104 ± 0.1 min–1 and 1.4 x 104 ± 0.2 min–1, respectively. The T. neapolitana AK-N and AK-C fragments also had detectable activity (0.09 x 104 ± 0.01 min–1) when mixed together. The rate observed for this fragment mixture was 17-fold lower than that of AKTn and 300-fold greater than the sum of the rates obtained for either of the fragments alone, indicating that these fragments cooperatively function in vitro. In contrast, a mixture containing equimolar levels of the B. subtilis AK fragments exhibited a rate (0.8 ± 0.2 min–1) that was <4-fold greater than the sum of the rates obtained for the fragments and >10 000-fold lower than either of the full-length AK homologs. These findings are consistent with those from bacterial selections showing that the split AKBs is not functional in vivo at 40°C.
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Secondary structure analysis
The disparity in the functions of the AK fragments could arise because the T. neapolitana fragments retain greater secondary structure and more readily self-associate. To establish the relative secondary structure for each AK fragment, far-UV CD spectra were obtained. Figure 6A shows the mean residue ellipticity for each AK-N over a range of wavelengths. Comparison of these spectra to a reference set of proteins using the K2D algorithm predicts that B. subtilis and T. neapolitana AK-N both contain low levels of helix, 16 and 12%, respectively (Andrade et al., 1993). This is much lower than the level of helix (69%) observed in the AKBs crystal structure for these fragments (Fig. 1), suggesting that both AK-N fragments are largely unstructured.
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Figure 6B shows the mean residue ellipticity for each AK-C fragment over a range of wavelengths. B. subtilis AK-C displays a strong negative peak at 203 nm, suggesting that this polypeptide is largely disordered. T. neapolitana AK-C, in contrast, has clear peaks at 208 and 222 nm, consistent with the presence of greater helical content. However, a comparison of this spectrum to a reference set of proteins using the K2D algorithm (Andrade et al., 1993
) suggests that its overall helix content (21%) of AK-C is still significantly lower than the 50% helix predicted for this fragment from the structure of other AK homologs (Fig. 1). To determine whether the T. neapolitana fragment pair acquires greater parent-like secondary structure upon mixing compared with the B. subtilis fragment pair, we investigated how the CD spectrum of each fragment pair changes upon mixing. Figure 7A shows the spectrum obtained at 40°C for a mixture containing B. subtilis AK-N and AK-C (10 µM each) and the spectrum calculated for a mixture using the spectra acquired with the individual fragments. These spectra are similar indicating that the AKBs fragments do not acquire increased parent-like structure when mixed at this concentration. In contrast, the spectrum obtained under similar conditions for a mixture of T. neapolitana fragments (10 µM) at 40°C exhibits a 17% decrease in ellipticity at 222 nm compared with the spectrum calculated from measurements of the individual fragments (Fig. 7). This indicates that the T. neapolitana fragments self associate upon mixing and acquire increased levels of parent-like secondary structure. However, the ellipticity observed for the mixture is less than that of AKTn at 222 nm (–15.2 x 10–3 deg·cm2/dmol), indicating that the split enzyme has less overall helical content than the full-length AK in the absence of substrates.
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Zinc content of AK fragments
AKBs and AKTn coordinate a zinc ion, and AKTn exhibits increased thermostability when zinc is bound (Vieille et al., 2003), suggesting that the differences in split protein structure and function could arise from changes in zinc stoichiometry. Whereas AKBs uses three cysteines and an aspartate in its AK-C fragment to coordinate zinc (Bae and Phillips, 2004), AKTn uses four cysteines found at structurally-related positions (Vieille et al., 2003). To quantify the relative zinc levels in each AK-C fragment, the purified polypeptides were mixed with a 40-fold excess of MMTS, a reagent that reacts specifically with sulfhydryl groups (Smith et al., 1975), and zinc release was monitored using PAR, a zinc chelator that exhibits increased absorbance at 500 nm upon zinc binding (Hunt et al., 1985). When MMTS was added to the T. neapolitana AK-C in the presence of excess PAR, the absorbance increased to a level that corresponded with 0.84 ± 0.02 equivalents of zinc released per AK-C. Similar experiments with B. subtilis AK-C revealed that this protein contains a reduced number of equivalents of zinc (0.62 ± 0.01).
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Discussion |
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We present evidence that the level of AK fragment complementation is directly proportional to the stability of the parents bisected: AKTn>AKBsQ16L/Q199R>AKBs. E. coli CV2T7 growth at 40°C was only complemented by AK fragments derived from the hyperthermophilic AKTn, which exhibits a mid-point for thermal denaturation (Tm) that is near the boiling point of water (Vieille et al., 2003
). In contrast, polypeptides derived from the mesophilic B. subtilis homolog with the lowest thermostability (Tm = 51–55°C) were non-functional under similar conditions, even though full-length AKBs displayed similar steady-state activity as AKTn at the temperature where growth selections were performed. Weak AKBs fragment complementation occurred on LB-agar plates when the Q16L and Q199R mutations were incorporated that increase the Tm for the full-length protein by 14°C (Counago et al., 2006). The growth of CV2T7 containing these fragments on LB-agar was slower than that of cells containing split AKTn, and they could not complement growth in liquid medium like AKTn.
The enhanced complementation observed when AKBs fragments were fused to associating IAAL-E3 and IAAL-K3 peptides suggests that split AK proteins can be used as an assay to report on protein–protein interactions in organisms where AK selections are available (Counago and Shamoo, 2005). In addition, these findings provide further evidence that parental thermostability influences the cooperative function of protein fragments. E. coli CV2T7 growth was poorest on LB-agar plates when cells harbored tagged AKBs fragment pairs that lacked mutations and those containing the Q199R mutation, which only increases the Tm for AKBs <2°C (Counago et al., 2006). In contrast, E. coli CV2T7 growth was strongest for tagged AKBs fragments containing the Q16L mutation, which is thought to be more thermostabilizing than Q199R (Counago et al., 2006).
Our findings are consistent with those from a study examining the folding and function of AK chimeras created by recombining mesophilic and thermophilic homologs (Bae and Phillips, 2006). With these chimeras, those having only their counterweight loops derived from residues in the mesophilic AK exhibited the smallest reduction in thermostability (Bae and Phillips, 2006). This is similar to our finding that peptide backbone cleavage within this region can yield a split AKTn that is functional like the full-length protein. In addition, protein thermostability was most strongly correlated with the fraction of the core residues inherited from the thermophilic parent in these chimeras (Bae and Phillips, 2006). These residues are found in both the AK-N and AK-C fragments, and of the AKBs fragments assayed, only the polypeptide containing the most thermostabilizing mutation (Q16L) functioned cooperatively with a T. neapolitana AK fragment.
Some hyperthermophilic enzymes are thought to achieve their thermostability through increased residual structure in their unfolded state (Robic et al., 2003). Our results provide evidence that this occurs when AK homologs having different thermostability are bisected at a single structurally-related position. Whereas the N-terminal fragments from AKBs and AKTn both had similar low levels of secondary structure, the C-terminal fragment derived from T. neapolitana AK exhibited greater parent-like helical content compared with the B. subtilis fragment. The AKTn C-terminal fragment also contained a higher fraction of bound zinc. Previous studies examining the role that zinc plays in regulating AK thermostability have shown that zinc removal reduces the Tm of Geobacillus stearothermophilus and T. neapolitana AK by 7.5°C and 6.2°C, respectively (Glaser et al., 1992; Vieille et al., 2003). This suggests that the reduced activity of the split AKBs at 40°C compared with the split AKTn could arise in part because AKBs has a lower amount of bound zinc. However, we were unable to directly assess the influence of zinc removal on the activity of the split AK homologs because the purified C-terminal fragments aggregated upon displacement of the zinc by MMTS.
Previous complementation studies with fibronectin type III domain and β-lactamases provided anecdotal evidence that protein thermostability and fragment complementation are correlated in single domain proteins. With the fibronectin type III domain, stabilizing mutations were identified using a yeast-two hybrid assay that improved complementation among fragments derived from a destabilized mutant (Dutta et al., 2005). When these mutations were incorporated into the full-length domain containing the same mutation, they all increased protein thermostability. In addition, the incorporation of a mutation shown to thermostabilize β-lactamases by 2.7 kcal/mol (Wang et al., 2002) led to a more sensitive protein fragment complementation assay (Galarneau et al., 2002). When lactamase fragments were fused to interacting proteins, they had increased catalytic activity. The results presented herein are the first to show that the cooperative function of protein fragments can be tuned by choosing naturally-occurring protein orthologs with different thermostabilities.
Protein evolution is driven by a variety of processes, including random mutation, intragenic recombination, domain insertion, and domain fission. Thermostability is known to influence a protein’s tolerance to amino acid substitutions created randomly and by recombination (Bloom et al., 2005; Bloom et al., 2006; Meyer et al., 2006; Besenmatter et al., 2007). Thermodynamic models have proposed that this arises because random substitutions produce similar average effects on the free energy of folding for proteins with the same topology (Bloom et al., 2005). Thus, when protein homologs incur similar mutations, the homolog with the lowest free energy of folding (i.e., greatest thermostability) is least likely to have its structure destabilized and its function altered. Our results suggest that a similar rationale can be used to explain the effects of backbone bisection on protein function. We hypothesize that upon fission of homologous proteins at structurally-related sites, fragmentation has similar destabilizing effects on protein homologs, and proteins with increased thermostability will on average be more likely to retain parent-like structure and function. Therefore, a quick way to create protein fragments with a range of cooperative functions is to select structurally-related parent proteins with a range of thermostabilities. Such an approach is expected to accelerate the directed evolution of protein fragments that cooperatively function (Ostermeier et al., 1999) for uses in proteomics (Michnick, 2003), drug discovery (Michnick et al., 2007), and synthetic biology (Giesecke et al., 2006).
One question raised by our observations is whether our split AKTn can complement an extremophile with a temperature-sensitive AK (Counago and Shamoo, 2005). Ongoing studies are evaluating how temperature affects AK fragment structure and function and examining whether a split AK could be used to generate protein fragment complementation assays for thermophiles (60–80°C) and hyperthermophiles (80°C) (Rothschild and Mancinelli, 2001). The development of such assays should facilitate the discovery of protein–protein interactions in microbes where classical two-hybrid systems have had limited success mapping genome-wide macromolecular interactions (Usui et al., 2005).
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This work was supported by the Robert A. Welch Foundation C-1614 (to J.J.S.) and the National Institutes of Health Biotechnology Training Grant 2T32-GM008362 (P.Q.N.).
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Abbreviations: AKBs, B. subtilis AK; AKEc, E. coli AK; AKTn, T. neapolitana AK; CD, circular dichroism; DTT, dithiothreitol; LB, luria broth; MMTS, methyl methanethiolsulfonate; PAR, 4-(2-pyridylazo) resorcinol; Tm, midpoint of thermal denaturation.
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TopAbstractIntroductionMaterials and methodsResultsDiscussionFundingAcknowledgementsReferences |
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We thank Yousif Shamoo for providing us with purified AKBs, Claire Vieille for providing pTNAK2:Km, and Corey J. Wilson and Kevin G. Hoff for critical comments.
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Andrade M.A., Chacon P., Merelo J.J., Moran F. Protein Eng. (1993) 6:383–390.
Bae E., Phillips G.N. Jr. J. Biol. Chem. (2004) 279:28202–28208.
Bae E., Phillips G.N. Jr. Proc. Natl Acad. Sci. USA (2006) 103:2132–2137.
Besenmatter W., Kast P., Hilvert D. Proteins (2007) 66:500–506.[CrossRef][Medline]
Bloom J.D., Silberg J.J., Wilke C.O., Drummond D.A., Adami C., Arnold F.H. Proc. Natl Acad. Sci. USA (2005) 102:606–611.
Bloom J.D., Labthavikul S.T., Otey C.R., Arnold F.H. Proc. Natl Acad. Sci. USA (2006) 103:5869–5874.
Cabantous S., Terwilliger T.C., Waldo G.S. Nat. Biotechnol. (2005) 23:102–107.[CrossRef][Web of Science][Medline]
Counago R., Shamoo Y. Extremophiles (2005) 9:135–144.[CrossRef][Medline]
Counago R., Chen S., Shamoo Y. Mol. Cell (2006) 22:441–449.[CrossRef][Web of Science][Medline]
de Prat Gay G., Ruiz-Sanz J., Fersht A.R. Biochemistry (1994) 33:7964–7970.[CrossRef][Web of Science][Medline]
Dutta S., Batori V., Koide A., Koide S. Protein Sci. (2005) 14:2838–2848.[CrossRef][Medline]
Fields S., Song O. Nature (1989) 340:245–246.[CrossRef][Medline]
Galarneau A., Primeau M., Trudeau L.E., Michnick S.W. Nat. Biotechnol. (2002) 20:619–622.[CrossRef][Web of Science][Medline]
Gegg C.V., Bowers K.E., Matthews C.R. Protein Sci. (1997) 6:1885–1892.[Web of Science][Medline]
Giesecke A.V., Fang R., Joung J.K. Mol. Syst. Biol (2006) 2:2006–2011.
Glaser P., Presecan E., Delepierre M., Surewicz W.K., Mantsch H.H., Barzu O., Gilles A.M. Biochemistry (1992) 31:3038–3043.[CrossRef][Web of Science][Medline]
Haase G.H., Brune M., Reinstein J., Pai E.F., Pingoud A., Wittinghofer A. J. Mol. Biol. (1989) 207:151–162.[CrossRef][Medline]
Hocker B., Beismann-Driemeyer S., Hettwer S., Lustig A., Sterner R. Nat. Struct. Biol. (2001) 8:32–36.[CrossRef][Web of Science][Medline]
Hunt J.B., Neece S.H., Ginsburg A. Anal. Biochem. (1985) 146:150–157.[CrossRef][Web of Science][Medline]
Johnsson N., Varshavsky A. Proc. Natl Acad. Sci. USA (1994) 91:10340–10344.
Kato I., Anfinsen C.B. J. Biol. Chem. (1969) 244:1004–1007.
Koradi R., Billeter M., Wuthrich K. J. Mol. Graph (1996) 14:51–55. 29–32.[CrossRef][Web of Science][Medline]
Linding R., Jensen L.J., Diella F., Bork P., Gibson T.J., Russell R.B. Structure (2003) 11:1453–1459.[Medline]
Litowski J.R., Hodges R.S. J. Biol. Chem. (2002) 277:37272–37279.
Magliery T.J., Wilson C.G., Pan W., Mishler D., Ghosh I., Hamilton A.D., Regan L. J. Am. Chem. Soc. (2005) 127:146–157.[CrossRef][Web of Science][Medline]
Meyer M.M., Hochrein L., Arnold F.H. Protein Eng. Des. Sel. (2006) 19:563–570.
Michnick S.W. Curr. Opin. Biotechnol. (2003) 14:610–617.[CrossRef][Medline]
Michnick S.W., Ear P.H., Manderson E.N., Remy I., Stefan E. Nat. Rev. Drug Discov. (2007) 6:569–582.[CrossRef][Web of Science][Medline]
Monnot M., Gilles A.M., Girons I.S., Michelson S., Barzu O., Fermandjian S. J. Biol. Chem. (1987) 262:2502–2506.
Muller C.W., Schlauderer G.J., Reinstein J., Schulz G.E. Structure (1996) 4:147–156.[Medline]
Nyfeler B., Michnick S.W., Hauri H.P. Proc. Natl Acad. Sci. USA (2005) 102:6350–6355.
Ostermeier M., Nixon A.E., Shim J.H., Benkovic S.J. Proc. Natl Acad. Sci. USA (1999) 96:3562–3567.
Pasek S., Risler J.L., Brezellec P. Bioinformatics (2006) 22:1418–1423.
Paulmurugan R., Umezawa Y., Gambhir S.S. Proc. Natl Acad. Sci. USA (2002) 99:15608–15613.
Pelletier J.N., Campbell-Valois F.X., Michnick S.W. Proc. Natl Acad. Sci. USA (1998) 95:12141–12146.
Remy I., Michnick S.W. Proc. Natl Acad. Sci. USA (1999) 96:5394–5399.
Remy I., Michnick S.W. Nat. Methods (2006) 3:977–979.[CrossRef][Web of Science][Medline]
Rial D.V., Ceccarelli E.A. Protein Expr. Purif. (2002) 25:503–507.[CrossRef][Medline]
Robic S., Guzman-Casado M., Sanchez-Ruiz J.M., Marqusee S. Proc. Natl Acad. Sci. USA (2003) 100:11345–11349.
Rothschild L.J., Mancinelli R.L. Nature (2001) 409:1092–1101.[CrossRef][Medline]
Saint Girons I., Gilles A.M., Margarita D., Michelson S., Monnot M., Fermandjian S., Danchin A., Barzu O. J. Biol. Chem. (1987) 262:622–629.
Schulz G.E., Muller C.W., Diederichs K. J. Mol. Biol. (1990) 213:627–630.[CrossRef][Web of Science][Medline]
Shiba K., Schimmel P. Proc. Natl Acad. Sci. USA (1992) 89:1880–1884.
Singh A., Mai D., Kumar A., Steyn A.J. Proc. Natl Acad. Sci. USA (2006) 103:11346–11351.
Smith D.J., Maggio E.T., Kenyon G.L. Biochemistry (1975) 14:766–771.[CrossRef][Web of Science][Medline]
Snel B., Bork P., Huynen M. Trends Genet. (2000) 16:9–11.[Web of Science][Medline]
Spotts J.M., Dolmetsch R.E., Greenberg M.E. Proc. Natl Acad. Sci. USA (2002) 99:15142–15147.
Usui K., et al. Genome Biol. (2005) 6:R98.[CrossRef][Medline]
Vieille C., Krishnamurthy H., Hyun H.H., Savchenko A., Yan H., Zeikus J.G. Biochem. J. (2003) 372:577–585.[CrossRef][Medline]
Wang X., Minasov G., Shoichet B.K. J. Mol. Biol. (2002) 320:85–95.[CrossRef][Web of Science][Medline]
Wilson C.G., Magliery T.J., Regan L. Nat. Methods (2004) 1:255–262.[CrossRef][Web of Science][Medline]
Wycuff D.R., Matthews K.S. Anal. Biochem. (2000) 277:67–73.[CrossRef][Web of Science][Medline]
Received November 14, 2007; revised January 12, 2008; accepted January 18, 2008.
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