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[18] In Vivo and In Vitro Analysis of the Rhodobacter sphaeroides Chemotaxis Signaling Complexes By STEVEN L. PORTER, GEORGE H. WADHAMS, and JUDITH P. ARMITAGE Abstract

This chapter describes both the in vivo and in vitro methods that have been successfully used to analyze the chemotaxis pathways of R. sphaeroides, showing that two operons each encode a complete chemosensory pathway with each forming into independent signaling clusters. The methods used range from in vitro analysis of the chemotaxis phosphorylation reactions to protein localization experiments. In vitro analysis using purified proteins shows a complex pattern of phosphotransfer. However, protein localization studies show that the R. sphaeroides chemotaxis proteins are organized into two distinct sensory clusters—one containing transmembrane receptors located at the cell poles and the other containing soluble chemoreceptors located in the cytoplasm. Signal outputs from both clusters are essential for chemotaxis. Each cluster has a dedicated chemotaxis histidine protein kinase (HPK), CheA. There are a total of eight chemotaxis response regulators in R. sphaeroides, six CheYs and two CheBs, and each CheA shows a different pattern of phosphotransfer to these response regulators. The spatial separation of homologous proteins may mean that reactions that happen in vitro do not occur in vivo, suggesting great care should be taken when extrapolating from purely in vitro data to cell physiology. The methods described in this chapter are not confined to the study of R. sphaeroides chemotaxis but are applicable to the study of complex two‐component systems in general. Introduction

Bacteria use a biased random swimming pattern to reach better environments for growth, moving either away from repellents or toward attractants. The chemosensory pathway of E. coli, described in detail in other chapters of this volume, is probably the best understood sensory system in biology. In E. coli, chemoeffectors bind to specific chemoreceptors clustered at the poles of the cell (Maddock and Shapiro, 1993). There are five different chemoreceptors responding to a limited number of effectors. A change in chemoeffector concentration is signaled across the membrane, regulating the activity of an associated histidine protein kinase dimer, CheA. A decrease in attractant binding to the receptor results in trans‐autophosphorylation of the METHODS IN ENZYMOLOGY, VOL. 423 Copyright 2007, Elsevier Inc. All rights reserved.

0076-6879/07 $35.00 DOI: 10.1016/S0076-6879(07)23018-6

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CheA dimer (Borkovich et al., 1989; Ninfa et al., 1991). Two response regulators compete for binding to CheA, CheY, a single domain response regulator, and CheB, a methyl esterase activated via a response regulator domain (Hess et al., 1988a,b). When phosphorylated, CheY‐P has reduced affinity for CheA but increased affinity for FliM on the flagellar motor, where it induces switching in the direction of motor rotation and, hence, a change in swimming direction (Welch et al., 1993). The activity of CheB increases about 100‐fold on phosphorylation (Anand et al., 1998). It serves, with the constitutive methyl transferase, CheR, to reset the signaling state of the chemoreceptors in a process termed adaptation (Lupas and Stock, 1989; Springer and Koshland, 1977). Signaling occurs in about 100 ms whereas adaptation takes about 1 s, bringing memory into the system. The rate of spontaneous CheY‐P dephosphorylation is increased by CheZ, to allow signal termination within the time course required for spatial gradient sensing (Lukat and Stock, 1993). For many years, it has been apparent that chemotaxis in the photoheterotrophic species Rhodobacter sphaeroides is somewhat different from that of E. coli. R. sphaeroides is a photoheterotrophic bacterium, a member of the alpha proteobacteria. Sequencing of the R. sphaeroides genome revealed three loci, each encoding a complete putative chemosensory pathway (Mackenzie et al., 2001). Our research over the past decade has centered on identifying why a bacterium with a single, unidirectional stop/start flagellum should apparently require three related pathways to control the stopping frequency of that flagellum (Armitage and Macnab, 1987). Two of these putative pathways have been shown to be essential for normal chemo (and photo) sensory behavior (Hamblin et al., 1997; Porter et al., 2002; Shah et al., 2000), although the level of their expression changes under different growth conditions (Martin et al., 2006; Shah et al., 2000). Our research has combined in vitro analysis of the phosphotransfer behavior of the different purified protein components with in vivo studies examining both the cellular localization of these proteins and the effects of mutating the genes encoding these proteins. The in vitro analysis has identified similarities and differences in phosphotransfer kinetics between components, and also the fact that some but not all components can cross‐talk, despite sequence similarities (Porter and Armitage, 2002, 2004; Porter et al., 2006). Although there is cross‐talk between some phosphotransfer proteins in the test‐tube, these proteins did not complement deletions of apparently equivalent proteins from the other loci in vivo. One possible reason for this became clear when we examined the localization of the proteins of two pathways; one pathway localizes to the cell poles with the transmembrane chemoreceptors and the other pathway localizes to a cytoplasmic midcell position with the soluble chemoreceptors (Martin et al., 2003; Wadhams et al., 2000, 2002, 2003).

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This chapter illustrates the techniques we used to identify the pattern of cross phosphorylation, and also highlights why relying on this alone may give a false model of the chemosensory pathways (Fig. 1). Combining the different approaches outlined here has identified not only what can phosphotransfer, but what phosphotransfer reactions probably do occur in vivo and where these interactions happen. The chemosensory pathway is just one of multiple two‐component pathways in bacteria, some species having over 100 putative pathways (Ashby, 2004; Galperin, 2006). Dissecting these interaction networks is a major challenge for bacteriology. The similarities among the different systems mean that the techniques described here are generally applicable to analyzing the phosphorylation and localization of two other component pathways. A CheA1

CheA2

CheA4

CheA1

CheA2

CheA3

B CFP-CheA2

C

YFP-CheA3

YFP-CheA4

CheA2

CheA4

CheA2

CheA3

FIG. 1. Reinterpretation of in vitro CheA phosphorylation reactions in R. sphaeroides after in vivo imaging. (A) Phosphorylation reactions that have been detected in vitro using purified proteins. (B) The localization of CheA2, CheA3, and CheA4. CheA1 does not appear to be expressed, so its localization could not be determined. (C) The auto‐/heterophosphorylation reactions that are likely to occur in vivo. These reactions are the same as those in panel (A) except that all reactions involving CheA1 have been removed because CheA1 is not expressed and is not required for chemotaxis; also, the phosphorylation of CheA2 by CheA4 has been removed because these proteins localize to different regions of the cell.

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In Vitro Analysis of Signaling by the Kinase Cluster

R. sphaeroides has loci that could encode the proteins for three different chemosensory pathways (Mackenzie et al., 2001). Sequence comparisons suggest there are genes for 6 putative CheY, 2 CheB, 3 CheR, 4 CheA, but no CheZ proteins. Some have high levels of homology while others show variations on the E. coli theme. For example, CheA1 and CheA2 are very similar to the E. coli CheA, but CheA3 and CheA4 lack some of the usual domains (Porter and Armitage, 2004). To identify whether the HPKs and RRs of the putative R. sphaeroides chemosensory pathway show similar phosphotransfer kinetics to the E. coli pathway and how the proteins might relate in a signaling pathway, they were all purified and their phosphorylation kinetics measured. Protein Purification In vitro analysis of the chemotaxis signaling pathway requires modest quantities of purified proteins. These can be produced by overexpressing the proteins in E. coli, followed by purification. Histidine protein kinases and response regulators have been successfully purified using His and GST tagging strategies (Jimenez‐Pearson et al., 2005; Porter and Armitage, 2002; Sourjik and Schmitt, 1998). A complete description of protein purification methodologies is beyond the scope of this chapter. Some HPKs are transmembrane proteins and may prove difficult to purify in sufficient quantity for analysis. The following protocol works for many of the cytoplasmic R. sphaeroides chemotaxis proteins when overexpressed using the His‐tagging pQE expression vectors (Qiagen) and may be a useful starting point for designing purification strategies for related proteins from other species (Porter and Armitage, 2002): 1. Transform the pQE‐based expression plasmid into the E. coli expression strain M15pREP4 (Qiagen). The transformed cells are resistant to both ampicillin (100 g/ml) and kanamycin (25 g/ml); both antibiotics should be used throughout the induction. 2. Grow a 25 ml overnight starter culture at 37
in Luria broth containing appropriate antibiotics. 3. Dilute overnight culture in 500 ml of 2YT medium containing appropriate antibiotics. Grow cells at 37
with shaking (225 rpm) to an OD600nm ¼ 0.8. 4. Add IPTG to the cultures to a final concentration of 100 g/ml. Incubate cultures for 20 h at 18
with shaking at 225 rpm. 5. Harvest cells by centrifugation at 6000g for 15 min. Resuspend the cell pellet in 30 ml of lysis buffer (10% glycerol, 50 mM Tris‐HCl, 150 mM

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NaCl, 10 mM imidazole, 1 mM DTT, pH 8). Freeze cells at 20
. For the R. sphaeroides chemotaxis proteins, frozen resuspended cell pellets can be stored for up to 6 months without reduction in yield of purified protein. 6. Thaw the resuspended cell pellet and lyze cells by sonication on ice. Cell lysis can be followed by microscopic examination. For the Vibracell sonicator (Sonics & Materials Incorporated), 6  20 s full power bursts are sufficient for full lysis. 7. Centrifuge the lysate at 35000g for 15 min to remove insoluble material and cell debris. 8. Pour 1 ml of Ni‐NTA Agarose slurry into a chromatography column (for example, catalog number 731–1550, Bio‐Rad). Allow the column to settle for at least 10 min. 9. Equilibrate the nickel column with lysis buffer. 10. Apply the cleared lysate to the equilibrated nickel column. 11. Wash the column with at least 60 ml of lysis buffer. 12. Elute the protein using lysis buffer containing 500 mM imidazole. Collect 1 ml fractions of the eluate. Protein usually elutes in the first three fractions. 13. Assay the fractions for protein using, for example, a Bradford assay (BioRad). 14. Pool the fractions containing significant quantities of protein and dialyze overnight against lysis buffer lacking imidazole. 15. Assess protein purity by SDS‐PAGE. Single‐step His‐tag purification is usually sufficient for phosphorylation assays. However, if significant levels of contaminating proteins are present, these can be removed by other standard protein purification techniques, for example, gel filtration or ion exchange chromatography. 16. Measure the protein concentration. Use a protein concentrator to increase the concentration if necessary (for typical phosphorylation/ phosphotransfer assays, protein concentrations in excess of 20 M are adequate). 17. Store the protein at 20
in single‐use aliquots. Although freezing does not affect the activity of the R. sphaeroides chemotaxis proteins (frozen proteins retain their activity for in excess of 1 year), it is possible that freezing may affect the activity of other proteins. Short‐term storage at 4
is an alternative. Measuring Kinase Activity Autokinase activity can be detected by incubating the putative histidine protein kinase with [ ‐32P] ATP for various time periods; this allows the kinase to autophosphorylate, which, depending on the kinase, can require

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time‐periods ranging from seconds to hours. The reaction mixtures are then analyzed on SDS‐PAGE gels. If autophosphorylation has occurred, then radioactive bands at a size corresponding to the protein of interest will be detectable by either autoradiography or phosphorimaging. If no radiolabeling of the kinase has occurred, it could be due to a lack of kinase activity under the assay conditions. Kinase activity is usually controlled by a sensory domain; the inclusion/exclusion of the stimulus sensed by the sensory domain in the assay mixture may be necessary to increase kinase activity to a detectable level. Another possible reason for an apparent lack of kinase activity is that the kinase is a hybrid kinase (contains both kinase and receiver domains). It is possible that the kinase autophosphorylates slowly but then rapidly phosphotransfers to the receiver domain, which then rapidly dephosphorylates; in such a scenario, no phosphoprotein would accumulate so it would appear that the protein has no kinase activity (Rasmussen et al., 2006). In such cases, there are two options: (1) Perform an ATPase assay on the protein (Ninfa et al., 1991). Even though no phosphoprotein accumulates, there will still be conversion of ATP to ADP if the putative hybrid kinase has kinase activity; (2) Inactivate the receiver domain—the phosphorylatable aspartate within the receiver domain can be mutated to alanine to prevent phosphotransfer to the receiver domain. Any phosphoryl groups should then remain on the histidine phosphorylation site of the hybrid kinase (Rasmussen et al., 2005). The following protocol has been used to analyze the autophosphorylation of R. sphaeroides CheA1 and CheA2 (Porter and Armitage, 2002); it should form a suitable starting protocol for any histidine protein kinase, which can then be optimized to the system under study. 1. Dilute the HPK to a final concentration of 5 M in TGMNKD buffer (final concentrations: 50 mM Tris HCl, 10% (v/v) glycerol, 5 mM MgCl2, 150 mM NaCl, 50 mM KCl, 1 mM DTT, pH 8.0). 2. Incubate the reaction mixtures at 20
for 1 h. 3. Add 2 mM [ ‐32P] ATP (specific activity 14.8 GBq mmol1; this specific activity is prepared by mixing unlabeled ATP with a commercially available [ ‐32P] labeled ATP of higher specific activity, e.g., 220 TBq mmol1). 4. Following addition of ATP, sample the reaction mixtures at intervals (15, 30, 90, 240, 480, 1800, and 3600 s are good starting points but these can be varied to suit the rate of the kinase) by removing a 10 l aliquot of the reaction mixture and mixing with 5 l of the quenching reagent (3 SDS‐ PAGE loading dye (7.5% (w/v) SDS, 90 mM EDTA, 37.5 mM Tris HCl, 37.5% glycerol, 3% (v/v) ‐mercaptoethanol, 0.05% (w/v) bromophenol blue, pH 6.8). Store the quenched samples on ice until all the samples have been collected.

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5. Heat the quenched samples to 65
for 30 s, then return to ice. Phosphorylated histidine and aspartate residues are heat labile. Do not boil samples prior to SDS/PAGE. 6. Load samples onto an SDS‐PAGE gel. To minimize hydrolysis of phosphoproteins, run gels at 500 V for as short a time period as possible at 4
(preferably less than 1 h). Use prechilled SDS‐PAGE running buffer and a refrigerating circulating water bath to cool the gel core during the electrophoresis run. 7. To keep radioactive waste contained, it is preferable not to run the dye front off the end of the gel. Following the electrophoresis run, cut the dye front off the gel and rinse the gel briefly in deionized water. Sandwich the gel between two sheets of OHP (overhead projector) acetate. 8. Serially dilute [ ‐32P] ATP (specific activity 14.8 GBq mmol1) in water to generate standards ranging in concentration from 0.05 to 5 M. Spot 10 l of each of these standards onto a strip of 3 MM Whatman paper. Allow these to dry and sandwich between two sheets of OHP acetate. 9. Expose the gel and the standard strip to a phosphorimaging screen or X‐ray film. Due to their linear responses, phosphorimaging screens are recommended for quantification purposes; a 30 min exposure time is sufficient for detecting CheA kinase activity. 10. Bands corresponding to phosphorylated protein should be visible on either the X‐ray film or the phosphorimage. To quantify the extent of phosphorylation, it is necessary to use image analysis software capable of outputting total pixel intensity for a defined region (e.g., ImageQuant TL, Amersham). Using this software, the regions containing the standards and the phosphoprotein bands are selected and the total pixel intensity measured. Following background subtraction, the total band intensity can be converted to amount of phosphorylated protein by comparison with the standard curve. 11. The autophosphorylation reaction should follow pseudofirst order rate kinetics. Plot the timecourse ([HPK‐P] versus time) and fit to the following pseudofirst order rate equation using a mathematical analysis package (e.g., Origin, Microcal): [HPK‐P] ¼ [HPK‐P]final (1ekobs*t) where kobs is the pseudofirst order rate constant, and [HPK‐P]final is the final (maximal) concentration of HPK‐P. Both kobs and [HPK‐P]final should be allowed to vary during the fitting procedure. 12. Repeat the entire experiment at different [ATP] ranging from 0.01 mM to 2 mM. This allows kobs to be determined for a range of ATP concentrations. Plot a graph of kobs vs [ATP]. Fit this curve to the following equation: kobs ¼ kcat[ATP]/(Km þ [ATP]) where kcat is the turnover rate for the HPK and Km is the Michaelis constant for ATP. Both kcat and Km should be allowed to vary during the fitting procedure.

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13. HPKs are functional only as dimers. At 5 M, CheAs are almost completely dimeric; other HPKs may be different. It is advisable to measure kcat under a range of different [HPK]; kcat will increase as [HPK] is raised because a greater fraction of the HPK present will be dimerized. However, once the HPK concentration is high enough that all of HPK is dimerized, then no further increases in kcat will be seen as the [HPK] is increased. This protocol is flexible and will need to be modified to take into account the HPK under study; the incubation temperature can be varied and the assay buffer can be modified to include/exclude ligands that affect kinase activity. Mg2þ is essential for all the activity of all known HPKs, so it should always be included in any assay buffer. Heterophosphorylation Reactions Typically, HPKs are homodimeric proteins in which the kinase domain of one monomer phosphorylates a histidine residue within the other monomer. This process is referred to as autophosphorylation. Heterophosphorylation occurs when the kinase domain of one protein phosphorylates a histidine residue within another non‐identical protein. A variation of the measuring kinase activity protocol was used to analyze heterophosphorylation between the atypical CheAs, CheA3 and CheA4, from R. sphaeroides. Both proteins are missing some of the domains normally found in CheAs and, using the previous protocol, it was shown that neither protein could autophosphorylate. However, when a mixture of CheA3 and CheA4 was analyzed using the protocol, CheA3 became phosphorylated, indicating that CheA4 is able to (hetero)phosphorylate CheA3 (Porter and Armitage, 2004). A further variation of the measuring kinase activity protocol was used to examine potential heterophosphorylation reactions among the other R. sphaeroides CheAs, for example, the phosphorylation of CheA1 by CheA2. However, this analysis was complicated by the ability of CheA1 and CheA2 to autophosphorylate. For this reason, a CheA1 mutant defective in autophosphorylation (containing the G501K mutation within its kinase domain) was used as the phosphoacceptor and a CheA2 mutant (H46Q) lacking a phosphorylatable histidine residue as the phosphodonor. Since neither CheA1(G501K) nor CheA2(H46Q) can autophosphorylate when incubated alone, the phosphorylation of CheA1(G501K) that was observed when these proteins were mixed can be assumed to be the result of the kinase domain of CheA2(H46Q) phosphorylating CheA1(G501K) (Porter and Armitage, 2004). Similar heterophosphorylation experiments on the other CheAs revealed the phosphorylation network shown in Fig. 1A.

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Phosphotransfer from HPKs to Response Regulators Phosphotransfer from a phosphorylated HPK to response regulators can be detected by mixing prephosphorylated 32P labeled HPK with the response regulators. If phosphotransfer occurs, then radioactive label will be transferred from the kinase to the response regulator; this can be followed using SDS‐PAGE followed by autoradiography or phosphorimaging. The prephosphorylation of the HPK is achieved by incubating the HPK with [ ‐32P] ATP for a period of time that allows almost complete phosphorylation. After this time, some investigators prefer to remove the unincorporated ATP from the reaction mixture prior to use in phosphotransfer assays; this prevents rephosphorylation of the HPK following phosphotransfer to response regulator and ensures that each phosphorylated kinase can only participate in a single phosphotransfer reaction. Alternatively, the unincorporated ATP can be left in the reaction mixture. However, this means that, following the initial phosphotransfer to the response regulators, the kinase can rephosphorylate and thus participate in further phosphotransfer reactions. If the ATP is left in the phosphotransfer reactions, then it is necessary to perform a control reaction in which the HPK is omitted from the reaction to demonstrate that the response regulator does not autophosphorylate. Some HPKs (e.g., NtrB) have phosphatase activity on their cognate response regulators. This phosphatase activity can prevent the accumulation of phosphorylated response regulator in the phosphotransfer assay. This may result in no phosphorylated response regulator being detected; however, providing that the excess [ ‐32P] ATP has been removed from the reaction, a reduction in the level of phosphorylated HPK should still be measurable if phosphotransfer has occurred. The following protocol was designed for the analysis of phosphotransfer among the four R. sphaeroides CheAs and the six CheYs, and should be adaptable to the study of any two‐component system: 1. Follow steps 1 through 3 of the measuring kinase activity protocol. 2. Following addition of [ ‐32P] ATP to the kinase, incubate the reaction mixture for a period of time that allows almost complete phosphorylation of the kinase (30 min is sufficient for CheA). 3. Optional: Remove the unincorporated ATP from the reaction using a centrifugal protein concentrator; the ATP will pass through the membrane, while the protein will be retained. The concentrated protein solution should be diluted with 1 ml of TGMNKD buffer and concentrated; repeat at least twice to ensure the removal of the ATP. Protein recovery from this procedure is usually less than 100%; therefore, it is necessary to determine the concentration of the recovered protein using, for example, a Bradford assay (BioRad).

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4. Immediately before addition of response regulator, remove a 10 l aliquot of the reaction mixture and quench by mixing with 5 l of the quenching reagent (3 SDS‐PAGE loading dye (7.5% (w/v) SDS, 90 mM EDTA, 37.5 mM Tris HCl, 37.5% glycerol, 3% (v/v) ‐mercaptoethanol, 0.05% (w/v) bromophenol blue, pH 6.8). Store the quenched samples on ice until all of the samples have been collected. 5. Add response regulator to a final concentration of 10 M (the concentration of kinase and response regulator can be varied in this assay if, for example, their cellular concentrations have been determined). 6. Following addition of the response regulator, sample the reaction mixtures at intervals (15, 30, 90, 240, 480, 1800, and 3600 s are good starting points but these can be varied to suit the rate of the phosphotransfer) by removing a 10 l aliquot of the reaction mixture and mixing with 5 l of the quenching reagent. Store the quenched samples on ice until all of the samples have been collected. 7. Process the samples and analyze them by SDS‐PAGE followed by autoradiography/phosphorimaging, as described in steps 5 through 10 of the measuring kinase activity protocol. If phosphotransfer has occurred, radioactive bands corresponding to the response regulators should be visible, which will be accompanied by a reduction in the amount of phosphorylated HPK (an example is shown in Fig. 2A). 8. Plot the concentration of HPK‐P and RR‐P at each time point to obtain the timecourse (see Fig. 2B for an example). This timecourse is determined by three processes: the rate of autophosphorylation of the HPK (if ATP was left in the reaction mixture), the rate of phosphotransfer from the HPK‐P to the RR, and the rate of dephosphorylation of the RR‐P. Estimation of the RR‐P dephosphorylation rate constant has been described in detail elsewhere (Porter and Armitage, 2002). Obtaining estimates for the rate constants describing the phosphotransfer reaction requires further experimentation, curve fitting, and mathematical modeling (Li et al., 1995; Stewart, 1997; Stewart et al., 2000).

Response Regulator Phosphatase Assays In addition to their intrinsic autophosphatase activity, the dephosphorylation of some response regulators is augmented by additional proteins, for example, CheZ, CheC, FliY, CheX, and RapA (Motaleb et al., 2005; Szurmant et al., 2004). The activity of these phosphatase proteins can be detected by performing parallel phosphotransfer experiments between the HPK and its cognate response regulator (as has been described; removal of the ATP following phosphorylation of the HPK is recommended) in the

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0

30

60

90

240

480

1920

CheA2-P

CheY4-P

B

6

[Protein-P] (mM)

5 4 3 2 1 0 0

100

200 300 Time (s)

400

500

FIG. 2. Example of a phosphotransfer timecourse. (A) CheA2 (5 M) was preincubated together with 0.5 mM [ ‐32P] ATP for 30 min. CheY4 (10 M) was then added to the reaction mixture (the final volume was 100 l). Ten microliter samples were removed at the indicated time‐points and quenched immediately by addition of 5 l of 3 SDS/EDTA loading dye. The quenched samples were analyzed by SDS‐PAGE and detected by phosphorimaging. (B) The experiment shown in Panel (A) was repeated three times and the radioactive band intensity quantified by phosphorimaging. Graphs show the mean concentrations of the phosphoproteins  the standard error of the mean.
, CheA2‐P; , CheY4‐P.

▪

presence and absence of the putative phosphatase. Phosphatase activity is suggested where there are reduced levels of RR‐P in the presence of the putative phosphatase. However, phosphatase activity is not the only explanation for reduced levels of RR‐P; an alternative possibility is that the putative phosphatase reduces the rate of phosphotransfer between the HPK and the RR. These two explanations are easily distinguished; if the rate of phosphotransfer were reduced, there would be an increase in HPK‐P in the reactions containing the putative phosphatase, whereas if the

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putative phosphatase were a phosphatase for the RR, then HPK‐P levels would either decrease or remain constant. Site‐Directed Mutagenesis of Kinases/RR to Confirm Role in Signaling While phenotypic analysis of a gene deletion/disruption mutant for a HPK or RR can demonstrate that the encoded protein is required for a particular cellular process, this gives no information on the role of phosphorylation in the control of the pathway. For example, CheA2 is essential for chemotaxis in R. sphaeroides (Martin et al., 2001), and protein localization studies have shown that CheA2 is necessary for the polar localization of CheW2, CheW3, and the transmembrane chemoreceptors (Wadhams et al., 2003, 2005). Is CheA2 essential for chemotaxis solely because of its role in localizing other proteins or is its phosphosignaling role also important? Deletion analysis could not answer this question. Instead, a CheA2 mutant was required that would support the localization of the other chemotaxis proteins but be defective in phosphorylation; one candidate was the phosphorylation site mutant CheA2(H46Q). This mutant was unable to support chemotaxis but resulted in correct localization of associated proteins, indicating that phosphosignaling from CheA2 is essential for chemotaxis (Porter and Armitage, 2004). 1. Identify putative phosphorylation sites by sequence alignment with other HPKs/RRs. 2. Mutate these putative phosphorylation sites. For response regulators, substitution of the phosphorylatable aspartate residue with alanine is recommended. Several cases of alternative site phosphorylation have been documented where the phosphorylatable aspartate was replaced with asparagine (Appleby and Bourret, 1999; Moore et al., 1993; Porter et al., 2006; Reyrat et al., 1994). 3. Purify the mutant HPKs/RRs and confirm using in vitro phosphorylation assays (described previously) that the phosphorylation site has been removed. 4. Compare the functionality of the mutant HPKs/RRs with their wild‐ type version in vivo. The most elegant way of doing this is to use a genomic replacement strategy, where the wild‐type gene is directly replaced with the mutant gene. If this is not possible in the organism of choice, then the ability of the wild‐type and mutant genes (expressed off plasmids at wild‐type levels) to complement a deletion strain can be compared. 5. If the wild‐type and mutant genes behave similarly, phosphorylation is not important for function of the HPKs/RRs. However, if there is a difference, this indicates that phosphorylation is important for function.

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Genomic Replacements with Fluorescent Protein Fusions for Studying Protein Localization

The in vitro analysis provided a very complex possible network of phosphotransfer reactions. We therefore examined the localization of each protein in the cell by fluorescently tagging each protein. This was carried out by replacing the wild‐type gene in the R. sphaeroides genome with a gene encoding a fusion protein. The gene was inserted in the normal operon position to ensure normal expression patterns. Whether to express a fusion protein from an inducible expression plasmid or by replacing the wild‐type gene in the genome with its fluorescent fusion version depends, to a certain extent, on the protein of interest and the organism being used. In general, expression from a plasmid is easier since it requires fewer genetic manipulations. However, it is imperative that the level of expression of the fusion protein is carefully checked to ensure that it is the same as that of the wild‐type protein, particularly in systems where there are large macromolecular complexes and stoichiometry might be important. Western blotting with an antibody raised to the wild‐type protein is the most common method for measuring expression levels. However, this only determines that the average expression level in a population of cells is similar to wild‐type. As the amount of inducer transported into each cell will vary, individual cells may express significantly more or less of the fusion than they would wild‐type protein. Expression of the fusion protein by direct replacement of the wild‐type gene in the genome requires the ability to make unmarked changes within the genome. The significant advantage of this technique is that the fluorescent fusion protein is expressed behind the native gene’s promoter elements and therefore should be expressed at the same levels and in the same location as the wild‐type protein. This can be especially important with genes whose expression may vary with growth conditions and for those usually expressed in stoichiometric quantities with other components of a system, for example, within an operon. However, when inserting fusion genes into operons, it is essential to ensure that the fusion protein does not interfere with any regulatory sequences for adjacent genes. As with expression from a plasmid, it is important to confirm wild‐type expression levels of the fusion protein by western blotting or a similar technique. It should also be confirmed that the expression of downstream genes in an operon has not been affected by the introduction of the fusion gene. There are numerous different fluorescent proteins available commercially and a detailed description of the various proteins and their properties is outside the scope of this chapter. In general, the proteins are derivatives of either GFP from the jellyfish Aequoria victoria or of coral reef proteins.

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These proteins are available in a wide range of spectral variants and some of them change colors when illuminated with light of a particular wavelength. The properties of different fluorescent proteins have been reviewed (Shaner et al., 2005). The fluorescent fusion can be to either the N‐ or C‐terminus of the protein and may involve the addition of flexible linkers between the fluorescent protein and the protein of interest to assist in the correct folding of the protein and thus retention of function. The choice of N‐ or C‐terminus for the fluorescent protein is often empirical, although if one terminus is known to be essential for function or interaction with other proteins, this terminus is often best avoided, at least initially. In either case, it is essential that the gene of interest and the gene encoding the fluorescent protein are in frame and there is no stop codon or start codon in the region linking the two proteins. The following describes the protocols used to replace genes with their corresponding fluorescent protein fusions in the genome of R. sphaeroides and methods for image acquisition and the subsequent analysis of these data. Fluorescent Protein Fusion Constructs for Integration into the Genome The first step in introducing an egfp tag into the R. sphaeroides genome involves generating the desired sequence within the suicide plasmid pK18mobsacB (Scha¨fer et al., 1994). The egfp sequence needs to be flanked by upstream and downstream sequences that are found within the R. sphaeroides genome, so that double homologous recombination will result in the introduction of egfp into the R. sphaeroides genome between these flanking sequences (Fig. 3). N‐Terminal Fusions In general, for an N‐terminal fusion a construct is generated that contains 500 bp immediately upstream of the gene fused to egfp, followed by the first 500 bp of the gene that must be in‐frame with the egfp (Fig. 4). The stop codon from egfp and the start codon for the gene should be excluded from the gene fusion. To ensure that the native Shine‐Dalgarno sequence is used to control expression of the fusion protein, overlap‐ extension PCR is used to position the start codon of egfp in exactly the same position as the start codon of the wild‐type gene, providing that this does not interfere with any stop codons or other important features of an upstream gene in the operon. If the stop codon from a preceding gene overlaps the start codon of the gene, it may be necessary to position the first codon of egfp a few codons into the existing gene. If the fusion is to a membrane protein, any membrane targeting sequence must be included before egfp for correct protein localization. As GFP folds as an autonomous

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5⬘

gfp

knR Suicide plasmid (pK18mobsacB derivative containing the fluorescent genefusion and flanking sequences) Introduced into R. sphaeroides by conjugation

3⬘

Genomic DNA of parental strain 3⬘

5⬘

Homologous recombination of suicide plasmid with genome. Integration selected for by plating on kanamycin

3⬘

5⬘

knR

3⬘

5⬘

sacB

Growth on kanamycin-free media, followed by plating on sucrose agar plates allows a second homologous recombination event to occur, excising the vector from the genome 3⬘

5⬘

5⬘ 3⬘ sacB knR

3⬘

5⬘

5⬘ 3⬘ sacB knR

Second recombination involved different regions to the first recombination

5⬘

3⬘

Wild-type gene replaced in the genome with the fluorescently tagged version

Second recombination involved the same regions as the first recombination

5⬘

3⬘

Parental strain

FIG. 3. Diagram outlining the steps involved in chromosomal gene replacement in R. sphaeroides.

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C

A

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UP

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407

F

2 3

A UP eGFP

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FIG. 4. Diagram showing the production of a construct for introducing the gene encoding an N‐terminal eGFP fusion into the R. sphaeroides genome. (1) Primers C and D are used to amplify the egfp gene. (2) Primers A and B are used to amplify the 500 bp region immediately upstream of the gene to be tagged. (3) Primers E and F are used to amplify the first 500 bp of the gene to be tagged. (4) Primers B and C are exactly complementary to one another, facilitating an overlap extension reaction between the products of reactions 1 and 2. The overlap extension product is amplified using primers A and D. (5) The products of PCRs 3 and 4 are ligated into the suicide vector pK18mobsacB to generate the tagging construct. Restriction sites within primers are shown in black; the restriction sites used in primers D and E must be compatible.

unit, extra amino acids prior to the GFP itself are fine, but the whole of the target protein (with the exception of the initiating methionine and any cleaved signal peptide) should always be included after the end of the GFP protein. The insert is ligated into a suicide vector, which integrates into the chromosome of the wild‐type strain replacing the wild‐type gene by double homologous recombination. Example Protocol 1. A 500 bp region immediately upstream of the gene of interest is amplified by PCR to include a suitable 50 restriction site. 2. egfp is amplified by PCR from pEGFP‐N1 (BD Biosciences) to include a suitable 30 restriction site and to exclude the egfp stop codon.

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3. The central primers (B & C; Fig. 4) are designed so the 30 end of the upstream fragment is complementary to the 50 end of the egfp fragment and vice versa. The two PCR products are combined and used as templates in overlap‐extension PCR with the 50 primer from the original upstream PCR and the 30 primer from the original egfp PCR. The product contains the region immediately upstream of the gene of interest fused to egfp with the start codon for egfp positioned exactly in the position of the start codon for the gene of interest. 4. This PCR product is cloned into the suicide vector pK18mobsacB. 5. The first 500 bp of the gene of interest is amplified by PCR (excluding its initial ATG) to include an upstream restriction site compatible with that on the 30 end of the overlap‐extension PCR product and a suitable downstream restriction site. This PCR product is cloned into the previous plasmid, generating a construct with the region upstream of the gene of interest fused to egfp and then 500 bp of the gene of interest. It is important that the egfp and the gene of interest are in‐frame. C‐Terminal Fusions When constructing a C‐terminal genomic fusion, similar principles apply as those for N‐terminal fusions; however, a restriction site can be included between the end of the gene of interest and the beginning of egfp. The basic construct for a C‐terminal fusion has 500 bp immediately upstream from the stop codon of the gene of interest, followed in‐frame by egfp and then the stop codon for the gene of interest and 500 bp immediately downstream of that stop codon. If the gene of interest is part of an operon, it is important that the Shine‐Dalgarno sequence or any other control elements for any downstream gene is not removed. In some cases, the start of a downstream gene overlaps the end of the gene of interest. The 30 region of the gene of interest can be repeated after egfp to ensure that potential control elements are retained. The insert is ligated into a suicide vector that integrates into the chromosome of the wild‐type strain, replacing the wild‐type gene by double homologous recombination. Confirm that expression of downstream genes in an operon is unaffected by the presence of the fusion using a technique such as western blotting. Example Protocol 1. A 500 bp fragment immediately preceding the stop codon of the gene of interest is amplified by PCR using primers that incorporate suitable restriction sites. 2. egfp without its initiating ATG is amplified by PCR using primers to introduce a restriction site at the 50 end compatible with the 30

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restriction site on the upstream fragment so that ligation results in the gene of interest and egfp being in‐frame. 3. A 500 bp fragment immediately downstream of and including the stop codon of the gene of interest is amplified by PCR using primers that include a 50 restriction site compatible with that on the 30 end of egfp. 4. The three fragments are ligated together into the suicide vector pK18mobsacB. Integration of Gene Fusions into the R. sphaeroides Genome The unmarked integration of genes into the R. sphaeroides genome is achieved by double homologous recombination (Fig. 3). Initially, the suicide plasmid pK18mobsacB integrates into the genome because of the homology between one of the flanking regions of the insert and the genomic DNA. The kanamycin resistance gene on the vector backbone allows selection of the recombinants. The cells are then grown in the absence of kanamycin selection and plated on sucrose‐containing medium. Cells retaining the vector backbone are lost since the sacB gene on the vector backbone converts sucrose to toxic levansucrose. Southern blotting and PCR following the second recombination determines whether the two recombination events have happened within the same flanking region, resulting in reversion to wild‐type, or in different flanking regions, resulting in the unmarked integration of the fusion gene into the genome. Protocol 1. Transform the pK18mobsacB construct into E. coli S17–1lpir. 2. Grow E. coli S17‐1 cells containing pK18mobsacB overnight at 37
in LB with kanamycin (25 g/ml). Grow Rhodobacter sphaeroides WS8N for 2 days at 30
in succinate media with nalidixic acid (25 g/ml). 3. On the day of the conjugation: Transfer 350 l of S17‐1 containing pK18mobsacB into 5 ml fresh LB with kanamycin. Grow at 37
with shaking until early log phase (faint cloudiness). 4. Harvest 1 ml of the S17‐1 cells containing pK18mobsacB at 6000 rpm. Harvest 1 ml of R. sphaeroides cells. 5. Wash pellets very gently in 1 ml LB. Spin again at 6000 rpm. 6. Resuspend pellets very gently in 100 l of LB. 7. Gently mix 10 l of S17‐1 containing pK18mobsacB and 100 l of R. sphaeroides. Transfer gently to a sterile filter disc on a dry LB plate (aged 1 day). Incubate at 30
overnight. 8. Pick up the filter with sterile forceps and transfer it to an Eppendorf tube containing 800 l LB. Vortex vigorously. Spread plate 100 l

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9. 10. 11.

12. 13.

14. 15.

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onto several LB agar plates containing nalidixic acid (25 g/ml) and kanamycin (25 g/ml). Incubate plates at 30
for at least 2 days. Pick a single colony into succinate media with nalidixic acid (25 g/ml) and grow for 2 days at 30
. Make serial dilutions into M22 minimal media and plate out 100 l of neat, 1/10, 1/100, and 1/1000 dilution onto sucrose plates (M22 with 10% sucrose and 2% agar). Incubate plates at 30
for 3 to 5 days. Replicate plate by picking single colonies from the sucrose plates with a sterile toothpick and making crosses on an LB with nalidixic acid (25 g/ml) and kanamycin (25 g/ml) plate and on a LB with nalidixic acid (25 g/ml) plate (48 colonies per pair of plates). Incubate plates at 30
for 3 to 5 days. Pick colonies from the LB with nalidixic acid plates that have not grown on the LB with nalidixic acid and kanamycin plates for genomic DNA extraction and further analysis.

Assessing the Functionality of the Fluorescent Protein Fusions

It is important to determine whether the fusion protein is functional and, if not, the level of reduction in functionality. This indicates probable correct localization and interaction. Comparison of the phenotype with a wild‐type and deletion mutant provides a straightforward measure of function. If there is no deletion strain, or deletion does not produce an obvious phenotype, alternative methods of assessing the accuracy of the localization of the fusion protein should be investigated. Immuno‐gold electron microscopy using antibodies to the wild‐type protein show the usual localization pattern of the protein (Martin et al., 2003; Wadhams et al., 2003). Alternatively, co‐localization studies with proteins whose fusions are functional and the ability to remove that localization on deletion of other proteins within the same system all help validate localization results (Wadhams et al., 2005). Image Acquisition and Data Analysis A detailed description of fluorescence microscope systems suitable for visualizing the sub‐cellular localization of fluorescent protein fusions has been reviewed elsewhere (Lichtman and Conchello, 2005). However, for most bacterial systems, a microscope with either phase contrast or DIC, a 100 oil immersion objective, and a mercury arc lamp or laser for excitation of the fluorescent protein are required. Sets of filters are also needed that are matched to the excitation and emission properties of the fluorophore being used. A cooled CCD camera is often used for higher resolution.

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For any semi‐quantitative data analysis it is important to ensure that the intensity of the excitation light and the exposure time are optimized to ensure that the fluorescence image is not saturating the camera’s pixels, but is not so dim that fluorescence is lost in any background noise. Usually, trying to fill the pixel wells to about 90% of their maximum intensity is a good starting point. It is also important that the wild‐type strain without any fluorescent fusion is imaged with the same microscope and camera settings on the same day as the strain containing the fluorescent fusion protein. This gives a background reading for cellular autofluorescence not attributable to the presence of the fluorescent protein fusion. If the objective of the experiment is simply to determine where within the cell the tagged protein is localized, then obtaining a bright field image showing the position of the cell bodies and a fluorescent image that can be superimposed on the first image will demonstrate protein localization (Fig. 1B). Many different software packages are available that will do this. The important principle is that any manipulation made to the images from the cells expressing the fluorescent protein fusion is also applied to images acquired with the same settings on the same day from cells without the fusion to ensure that observed fluorescence is directly attributable to the fusion protein. Summary

Protein purification and analysis of phosphotransfer rates provide an accurate quantitative measure of the kinetics of phosphotransfer reactions. However, while the reactions tell you what can interact and phosphotransfer, they do not reveal which HPKs and RRs do really interact. If there are multiple homologues in a system, a combination of biochemical and microscopic approaches can provide a realistic model of the sensory network. For example, the CheAs of R. sphaeroides show complex heterophosphorylation patterns in vitro (Fig. 1A), but upon analysis of the cheA deletion phenotypes and the cellular localization of the CheAs (Fig. 1B), the story became much simpler (Fig. 1C). There are two separate signaling clusters; although CheA4 can phosphorylate CheA2 in vitro, CheA2 is localized to the cell poles while CheA4 is localized to the cytoplasmic chemotaxis cluster. This spatial separation of CheA2 and CheA4 would therefore make it unlikely that CheA4 phosphorylates CheA2 in vivo. References Anand, G. S., Goudreau, P. N., and Stock, A. M. (1998). Activation of methylesterase CheB: Evidence of a dual role for the regulatory domain. Biochemistry 37, 14038–14047. Appleby, J. L., and Bourret, R. B. (1999). Activation of CheY mutant D57N by phosphorylation at an alternative site, Ser‐56. Mol. Microbiol. 34, 915–925.

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Porter, S. L., and Armitage, J. P. (2004). Chemotaxis in Rhodobacter sphaeroides requires an atypical histidine protein kinase. J. Biol. Chem. 279, 54573–54580. Porter, S. L., Warren, A. V., Martin, A. C., and Armitage, J. P. (2002). The third chemotaxis locus of Rhodobacter sphaeroides is essential for chemotaxis. Mol. Microbiol. 46, 1081–1094. Porter, S. L., Wadhams, G. H., Martin, A. C., Byles, E. D., Lancaster, D. E., and Armitage, J. P. (2006). The CheYs of Rhodobacter sphaeroides. J. Biol. Chem. 281, 32694–32704. Rasmussen, A. A., Porter, S. L., Armitage, J. P., and Sogaard‐Andersen, L. (2005). Coupling of multicellular morphogenesis and cellular differentiation by an unusual hybrid histidine protein kinase in Myxococcus xanthus. Mol. Microbiol. 56, 1358–1372. Rasmussen, A. A., Wegener‐Feldbrugge, S., Porter, S. L., Armitage, J. P., and Sogaard‐ Andersen, L. (2006). Four signaling domains in the hybrid histidine protein kinase RodK of Myxococcus xanthus are required for activity. Mol. Microbiol. 60, 525–534. Reyrat, J. M., David, M., Batut, J., and Boistard, P. (1994). FixL of Rhizobium meliloti enhances the transcriptional activity of a mutant FixJD54N protein by phosphorylation of an alternate residue. J. Bacteriol. 176, 1969–1976. Scha¨fer, A., Tauch, A., Ja¨ger, W., Kalinowski, J., Thierbach, G., and Pu¨hler, A. (1994). Small mobilizable multipurpose cloning vectors derived from the Escherichia coli plasmids pK18 and pK19—Selection of defined deletions in the chromosome of Corynebacterium glutamicum. Gene 145, 69–73. Shah, D. S. H., Porter, S. L., Martin, A. C., Hamblin, P. A., and Armitage, J. P. (2000). Fine tuning bacterial chemotaxis: Analysis of Rhodobacter sphaeroides behavior under aerobic and anaerobic conditions by mutation of the major chemotaxis operons and cheY genes. EMBO J. 19, 4601–4613. Shaner, N. C., Steinbach, P. A., and Tsien, R. Y. (2005). A guide to choosing fluorescent proteins. Nat. Meth. 2, 905–909. Sourjik, V., and Schmitt, R. (1998). Phosphotransfer between CheA, CheY1, and CheY2 in the chemotaxis signal transduction chain of Rhizobium meliloti. Biochemistry 37, 2327–2335. Springer, W. R., and Koshland, D. E., Jr. (1977). Identification of a protein methyltransferase as the cheR gene product in the bacterial sensing system. Proc. Natl. Acad. Sci. USA 74, 533–537. Stewart, R. C. (1997). Kinetic characterization of phosphotransfer between CheA and CheY in the bacterial chemotaxis signal transduction pathway. Biochemistry 36, 2030–2040. Stewart, R. C., Jahreis, K., and Parkinson, J. S. (2000). Rapid phosphotransfer to CheY from a CheA protein lacking the CheY‐binding domain. Biochemistry 39, 13157–13165. Szurmant, H., Muff, T. J., and Ordal, G. W. (2004). Bacillus subtilis CheC and FliY are members of a novel class of CheY‐P‐hydrolyzing proteins in the chemotactic signal transduction cascade. J. Biol. Chem. 279, 21787–21792. Wadhams, G. H., Martin, A. C., and Armitage, J. P. (2000). Identification and localization of a methyl‐accepting chemotaxis protein in Rhodobacter sphaeroides. Mol. Microbiol. 36, 1222–1233. Wadhams, G. H., Martin, A. C., Porter, S. L., Maddock, J. R., Mantotta, J. C., King, H. M., and Armitage, J. P. (2002). TlpC, a novel chemotaxis protein in Rhodobacter sphaeroides, localizes to a discrete region in the cytoplasm. Mol. Microbiol. 46, 1211–1221. Wadhams, G. H., Martin, A. C., Warren, A. V., and Armitage, J. P. (2005). Requirements for chemotaxis protein localization in Rhodobacter sphaeroides. Mol. Microbiol. 58, 895–902. Wadhams, G. H., Warren, A. V., Martin, A. C., and Armitage, J. P. (2003). Targeting of two signal transduction pathways to different regions of the bacterial cell. Mol. Microbiol. 50, 763–770. Welch, M., Oosawa, K., Aizawa, S.‐I., and Eisenbach, M. (1993). Phosphorylation‐dependent binding of a signal molecule to the flagellar switch of bacteria. Proc. Natl. Acad. Sci. USA 90, 8787–8791.