.maamuwmm s. :5. I a u - ,g.I~n-W‘o 3 Jr - r. . . Itchy!“ .v I}? ,1; u i... T... [5.91.3.1 3.1.. .8..I...lsi..fiwutfl, 1t .12... "92...th .l . i 1: iii... 2 4%E-....,..3.5.....a -fl............s.....u,.z... a... x 3 . .. _ 13.33! ‘21}:- ialxxll5 (.55.!!- v liltixh £75 .\ ) 53!. mi“; ' LIBDARY 10f / MiGhanr. State afl_UhyeEMy This is to certify that the dissertation entitled A NOVEL MECHANISM OF C-SIGNAL-DEPENDENT GENE REGULATION DURING MYXOCOCCUS XANT HUS DEVELOPMENT presented by SHEENU MI'I'TAL has been accepted towards fulfillment of the requirements for the PhD. degree in Cell and Molecular Biology 5? w: nK/vociu, Major Professor’s Signature 8/17/08 I I Date MS U is an Affirmative Action/Equal Opportunity Institution PLACE IN RETURN BOX to remove this checkout from your record. To AVOID FINES return on or before date due. MAY BE RECALLED with earlier due date if requested. DAJEDUE DAIEDUE DAIEDUE 5/08 K‘IPrqlAcc8Pres/ClRC/DateDue.nndd A NOVEL MECHANISM OF C-SIGNAL-DEPENDENT GENE REGULATION DURING M YXOCOC CUS XANTH US DEVELOPMENT By SHEENU MITTAL A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Cell and Molecular Biology 2008 ABSTRACT A NOVEL MECHANISM OF C-SIGNAL-DEPENDENT GENE REGULATION DURING M YXOC 0C C US XANT H US DEVELOPMENT By Sheenu Mittal The Gram-negative soil bacterium, Myxococcus xanthus, undergoes a multicellular developmental process upon nutrient limitation. Intercellular signals expressed at different times during the developmental process coordinate cell movements and regulate spatiotemporal gene expression. C-signal, a short-range signal that may require cell-cell contact, is essential for cells to move into aggregates and differentiate into spores within the mound-shaped fruiting bodies. To understand gene regulation in response to C-signaling, the control of two C-signal- dependent transcription units, whose products play a role in aggregation and sporulation, fmgA and fingBC, was studied. MrpC2 was discovered as a direct activator of fmgA and fingBC expression. MrpC2, an N-terminally truncated form of the protein, MrpC, is a development-specific transcription factor. MrpC functions as an antitoxin to mediate programmed cell death of about 80% of the cells during development. Gel-shift assays showed that MrpC2 binds to sequences important for fingA and fmgBC expression. Chromatin immunoprecipitation assays demonstrated association of MrpC and/or MrpC2 with the fmgA and fmgBC promoter regions in vivo. Previously, FruA, a transcription factor believed to be phosphorylated in response to C- signaling, was shown to be essential for fingA expression. MrpC2 binds to a sequence adjacent to the binding site of FruA in the fmgA promoter region. FruA was found to be essential for association of MrpC and/ or MrpC2 with the fmgA promoter region in vivo. DNase I footprinting revealed cooperative binding of FruA and MrpC2 to fingA promoter region DNA, and the combination of proteins enhanced formation of shifted complexes with fmgA DNA, unless the DNA had a mutation in the FruA- or MrpC2-binding site. This is a novel mechanism of gene regulation since cooperative binding of a response regulator (FruA) and another transcription factor (MrpC2) has not been observed previously. Expression of the fmgBC operon was found to be regulated by a similar mechanism as fingA. fmgB—lacZ expression depended absolutely on FruA. Association of FruA with the fmgBC promoter region was observed in vivo and in vitro. FruA bound to a key cis- regulatory sequence in the fmgBC promoter region in vitro, although it bound downstream of MrpC2, as opposed to binding upstream of MrpC2 in the fingA promoter region. Nevertheless, similar to fingA gene, FruA was required for in vivo occupancy of the fmgBC promoter region by MrpC and/or MrpC2. Also, FruA and MrpC2 appeared to bind cooperatively to fmgBC promoter region DNA in vitro. The results with fmgA and fmgBC (and preliminary results with two other C-signal- dependent genes) suggest that combinatorial regulation by FruA and MrpC2 is a conserved mechanism during M. xanthus development. This mechanism is proposed to monitor C-signaling via FruA phosphorylation, and persistent starvation via the MrpC2 level, resulting in sporulation of some cells and programmed cell death of others. To my parents, Rajendra and Rekha Mittal, and my husband, Shireesh Srivastava, without them I could not have accomplished this. iv ACKNOWLEDGMENTS I thank God almighty for what I am, for my parents, my husband and my family and friends. I would like to express my heartfelt thanks to my mentor, Dr. Lee Kroos, who helped me to develop into a good scientist. He had been very supportive in every possible way, throughout my graduate studies. I thank him for his patience and belief in me when things did not seem to work at all. His constant guidance and encouragement helped me finish my studies productively. I am grateful to my committee members, Dr. David Amosti, Dr. Zachary Burton, Dr. Cindy Arvidson, and Dr. Vincent Young for their very helpful suggestions towards my research. I thank them for taking time to attend the meetings and provide their perspectives which helped me understand the broader implications of my research. I would like to thank Kroos lab members for the scientific and friendly environment in the lab. Dr. Ruanbao Zhou was always there to answer my queries related to the experiments. I would like to express my gratitude to the past Kroos lab members: Dr. Deborah Himes, Dr. Pooma Viswanathan and Dr. Srinivasan Durairaj for their help and advice with regards to my research projects. Thank you also to the other members of the Kroos lab (past and present): Paul Himes, Christina Cusumano, Paul Luethy, Dr. Daisuke Imamura, Dr. Lijuan Wang, Dr. Bin Chen, Dr. Mark Robinson and Jun-seok Lee for insightful discussions. The confidence and unwavering support of my family and friends made it possible for me to come through all the obstacles faced during research. I am indebted to my parents for who I am. I thank them for the hardships and sacrifices they made to support my education and to provide the best to me and my siblings. I am especially thankful to my husband, Shireesh, for his never dying faith in me. I owe him this moment. He is a constant source of scientific and general knowledge. Despite of our varied research areas, we have always been able to understand each other’s work and provide helpful ideas. Shireesh has an amazing perspective towards life and I continue growing into a better individual with him. I would also like to thank my parents-in-law, Shishir and Kamlesh Srivastava, for their continuous encouragement and moral support. I thank my sisters, Anju and Meenu, my brother, Virendra, and rest of my family for their love and support. I would also like to thank my friends, Neelam and Trupti, for being fun to be with and for being helpful when my husband left for post-doctoral research a year and a half ago. Last but not the least, I thank the Cell and Molecular Biology program, the Van Andel Research Institute (VARI), the College of Natural Sciences, the Office of International Student and Scholars, and the Graduate School for the financial support in the form of VARI Graduate Fellowship, Dissertation Completion Fellowship and Travel Fellowships. Their contributions helped me present my research at the national and international levels. vi TABLE OF CONTENTS LIST OF TABLES .................................................................................. i x LIST OF FIGURES ................................................................................ x LIST OF ABBREVIATIONS .................................................................. xii CHAPTER 1: Introduction Motility ...................................................................................... 2 Signal transduction ............................................................................ 4 Sigma factors ............................................................................... 4 Enhancer-binding proteins ................................................................ 6 Fruiting body formation ................................................................... 7 Intercellular signaling during development................................................8 A-srgnalrng ..9 B-srgnalrnglO ..12 E-signaling ................................................................................... 12 C-srgnalrngl3 The C-signal transduction pathway ...................................................... 15 C-signal-dependent gene expression ..................................................... 19 CHAPTER 2: Cooperative binding of a response regulator and a bifunctional transcription factor/antitoxin controls gene expression and cell fate during Myxococcus xanthus development Abstract ...................................................................................... 26 Introduction ................................................................................. 27 Materials and Methods ..................................................................... 32 Results An insertion in fingA delays aggregation .......................................... 38 MrpC2 binds to a cis-regulatory sequence in the fmgA promoter region ................................................................................. 4O FruA is required for association of MrpC and/or MrpC2 with the fmgA promoter reggion in vivo ........................................................... 47 Cooperative binding of MrpC2 and FruA to the fmgA promoter region ................................................................................. 48 Discussion .................................................................................... 62 CHAPTER 3: Combinatorial regulation by a novel arrangement of FruA and MrpC2 transcription factors during Myxococcus xanthus development. Abstract ...................................................................................... 71 Introduction ................................................................................. 73 Materials and Methods ........................................................................ 78 Results An insertion in fingC reduces spore formation ................................... 83 vii MrpC2 binds to a key cis-regulatory sequence in the fmgBC promoter region ................................................................................. 84 MrpC and/or MrpC2 associates with the fmgBC promoter region in vivo and this depends on FruA ................................................................ 9O FruA associates with the fmgBC promoter region in vivo and governs expression. . . . . .............................................................................................. 92 The FruA DNA-binding domain binds to a key cis-regulatory sequence in the fmgBC promoter region ........................................................ 94 Enhanced complex formation in the presence of FruA-His6 and I-Iislo- MrpC2 ................................................................................ 94 Discussion ................................................................................. 102 CHAPTER 4: Conclusions and future directions Conclusions ................................................................................ 1 10 Future directions .......................................................................... 112 REFERENCES ..................................................................................... 1 18 viii LIST OF TABLES Table 2.1 Bacterial strains and plasmids used in this study ................................... 36 Table 3.1 Bacterial strains and plasmids used in this study .................................. 81 ix LIST OF FIGURES Figure 1.1 Regulatory network during M. xanthus development................................16 Figure 2.1 Model of the M. xanthus regulatory network and scanning electron micrographs of fruiting body development..........................................................39 Figure 2.2 Effects of mutations on fmgA promoter activity in vivo and on DNA binding in vitro ................................................................................................... 41 Figure 2. 3A SDS- PAGE of protein purified from the AS fraction using fingA DNA ( 101 to +25)... ... .... ...43 Figure 2.33 EMSAs with 32P-labeledfmgA DNA (6 nM) spanning from -101 to +25 and proteins in the AS fraction or the APP .............................................................................. 43 Figure 2.4 Anti-MrpC antibodies supershift the complex formed by fingA promoter region DNA and the AS fraction ................................................................... 44 Figure 2.5 Comparison of purified Hislo— MrpC2 and the AS fraction for binding to the fingA promoter region” . ...46 Figure 2.6 FruA is required for association of MrpC and/or MrpC2 with the fingA promoter region in vivo ............................................................................ 49 Figure 2.7A The combination of FruA- Hi56 and Hism-MrpC2 enhances complex formation. .... ... ... ...53 Figure 2.7B The combination of FruA-DBD-Hi38 and Hism-MrpCZ does not enhance complex formation .................................................................................. 53 Figure 2.8A DNase I footprinting shows cooperative binding of MrpC2 and FruA to the fingA promoter region (top strand labeled) ...................................................... 55 Figure 2.8B DNase I footprinting shows cooperative binding of MrpC2 and FruA to the fingA promoter region (bottom strand labeled) ................................................. 56 Figure 2.8C Summary of protected and hypersensitive sites .................................. 56 Figure 2.9 Enhancement of shifted complex formation depends on binding sites for both FruA and MrpC2 .................................................................................... 57 Figure 2.10A Supershift assay with an increasing amount of anti-MrpC antibodies (0.2, 0.3, 0.5 or 0.6 uM) as indicated ..................................................................... 61 Figure 2.10B Supershift assay with an increasing amount of anti-FruA antibodies (7 or 8 uM) as indicated .................................................................................... 61 Figure 3.1 Effects of mutations on fingBC promoter activity in vivo and on DNA binding in vitro ................................................................................................................................ 86 Figure 3.2A SDS-PAGE of protein purified from the AS fraction using fingBC DNA (- 104 to -29) ............................................................................................. 87 Figure 3.2B EMSAs with 32P-labeled fmgBC DNA (12 nM) spanning from -104 to -29 and proteins in the AS fraction or the APP ........................................................................ 87 Figure 3.3 Comparison of purified I-Iislo—MrpCZ and the AS fraction for binding to the fingBC promoter region ............................................................................. 89 Figure 3.4 Association of MrpC and/or MrpC2 with the fingBC promoter region during development of wild-type and fruA mutant cells ............................................................... 92 Figure 3.5 Association of FruA with the fingBC promoter region in vivo .................... 94 Figure 3.6 Developmental expression from fingB-lacZ ........................................ 96 Figure 3. 7 Effects of mutations on binding of FruA-DBD- Hiss to fingBC promoter region Figure 3.8A Shifted complex formation with Hislo-MrpC2 and full-length FruA-His6, and the effect of mutations ............................................................................ 100 Figure 3.8B Shifted complex formation with Hism-MrpCZ and FruA-DBD- HISg .................................................................................................................................. 100 xi 10. ll. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. LIST OF ABBREVIATIONS . ATP — adenosine triphosphate bp — base pair cGMP — cyclic guanosine monophosphate CTT — casitone—tris media DNA — deoxyribonucleic acid dNTP — deoxynucleotide triphosphate GTP - guanosine triphosphate His — histidine h — hour IgG — immunoglobulin G kb —- kilobase kDa — kilodalton Km — kanamycin M — molar Mb — megabase mM — millimolar min -— minute m1 — millilitre 1.1g — microgram ul - microlitre uM — micromolar NAD — nicotinamide adenine dinucleotide xii 23. ng — nanogram 24. nM - nanomolar 25. NTP -— nucleotide triphosphate 26. PAGE - pol yacrylamide gel electrophoresis 27. PCR — polymerase chain reaction 28. (p)ppGpp — guanosine (penta) tetraphosphate 29. RNA — ribonucleic acid 30. SDS - sodium dodecyl sulfate 31. TPM - tris-phosphate-magnesium media xiii Chapter 1: Introduction INTRODUCTION The myxobacteria were discovered in the nineteenth century and are Gram-negative bacteria that display a social lifestyle. Based on 168 ribosomal RNA sequence data, the myxobacteria belong to the delta-subgroup of the Proteobacteria (Shimkets and Woese 1992). They feed on organic matter in the soil, as well as on other bacteria and protozoa. When feeding, they travel like packs of wolves, secreting enzymes to degrade and hydrolyze food sources (Rosenberg et al. 1977). Among the myxobacteria, Myxococcus xanthus has been the best-studied. The sequence of M. xanthus strain DKl622 revealed a single circular chromosome of 9.14 Mb with 7,388 putative coding sequences (Goldman et a1. 2006). Gene duplication has been proposed to contribute to the large size of the M. xanthus genome. M. xanthus is a proficient producer of secondary metabolites including antimicrobials, polyketides (PKs), nonribosomally-made peptides (NRPs), and carotenoids (Goldman et al. 2006). Around 9% of the M. xanthus genome encodes enzymes involved in secondary metabolism. Motility M. xanthus is a rod-shaped, top-soil dwelling bacterium that has been isolated from many sites around the world. It lacks flagella, but moves by gliding on solid surfaces. M. xanthus employs two distinct motility systems for gliding; the A (adventurous) and S (social) systems (Hodgkin and Kaiser 1979a, Hodgkin and Kaiser 1979b). Lone cell movements have been attributed to A-motility and group movements to S-motility. There are currently two models to explain the A-motility mechanism. These models differ from one another in terms of the location of the gliding motor in a cell. According to the first model, there are small nozzle-like structures present at both poles of the cell and slime secretion from one end provides the required force to push cells forward (Wolgemuth et al. 2002). These nozzle-like structures have been proposed to be analogous to the nozzles that mediate gliding motility via slime secretion in cyanobacten'a (Hoiczyk and Baumeister 1998). According to the second model, the gliding motor is not localized at the poles, but instead it is distributed along the cell body (Sun et al. 1999, Mignot et al. 2007, Sliusarenko et al. 2007). According to this mode], membrane-anchored adhesion complexes exert force on the cytoskeleton to move the cell forward by pushing it against the substratum (Mignot et al. 2007). S-motility requires type IV pili, lipopolysacchan'de O—antigen, extracellular polysaccharide (EPS) or fibrils and FrzS (Li et al. 2003, Mignot et al. 2005). S-motility results from retraction of the polar pili, pulling the cell forward (Li et al. 2003). This retraction mechanism in M. xanthus is similar to that seen in Neisseria gonorrhoeae and Pseudomonas aeruginosa (Wolfgang et al. 1998, Skerker and Berg 2001). The frequency of cellular reversals during motility is controlled by genes of the Frz (Frizzy) chemosensory pathway in M. xanthus (Blackhart and Zusman 1985). The Frz pathway is similar to the Che (Chemotaxis) system found in enteric bacteria like E. coli. The Frz pathway regulates cell reversals, which is essential for directed cell movements during M. xanthus development. Elasticotaxis is another sensory mechanism found in M. xanthus. Elasticotaxis has been described as an effect that results in an asymmetrically elongated swarm on a compressed agar surface (Stanier). A-motility has been shown to play a role in elasticotaxis (Fontes and Kaiser 1999). Signal transduction M. xanthus harbors a vast number of proteins similar to those that typically form bacterial two-component systems (TCSs) including 137 His protein kinases (HPKs) and 118 response regulators (RRs) (Goldman et al. 2006). Two-component His/Asp systems are the most common bacterial signal transduction systems. They contain a His kinase which senses a signal via its N-terminal input domain, resulting in autophosphorylation of a His residue in the C-terminal transmitter domain. The phosphoryl group from the His residue is then transferred to a conserved Asp residue in the receiver domain of the RR. This results in the activation of the C-terminal effector domain of the RR. Most RRs contain a DNA-binding motif in the effector domain and when activated via phosphorylation, bind to DNA to activate gene expression. M. xanthus employs His-Asp signal transduction systems in a variety of processes including motility, development, pili biogenesis, heat shock, and osmotic tolerance (Whitworth 2008). The genome sequence also revealed an abundance of 102 eukaryotic-like protein Ser/T hr kinases (PSTKs) in M. xanthus (Goldman et al. 2006). The first bacterial PSTK was identified in M. xanthus (Munoz- Dorado et al. 1991) and PSTKs were subsequently discovered in many other bacteria (Av-Gay and Everett 2000, Wang et al. 2002, Petrickova and Petricek 2003). M. xanthus utilizes both TCSs and PSTKs to couple gene expression to continuously changing environmental cues. Sigma factors Additional mechanisms for coupling external stimuli with coherent cellular responses are provided by 38 putative extra-cytoplasmic function (ECF) sigma factors present in the M. xanthus genome (Goldman et al. 2006). ECF sigmas belong to the 0'70 family and typically respond to external stimuli to affect expression of genes whose products are involved in functions of the cell envelope and periplasrnic space (Helmann 2002). Of the 38 putative ECF sigmas, only three are well-characterized. CarQ regulates light-induced synthesis of carotenoids (Gorham et al. 1996). RpoEl plays a role in motility during vegetative growth and development (Ward et al. 1998). chA regulates expression of an early developmental gene (Kroos and Inouye 2008). Our group has created insertion mutants of each of the remaining 35 putative ECF sigma genes, and the mutants are being characterized for defects in motility, development, and secondary metabolite production, in collaboration with other groups to gain insight into the role of ECF sigma factors (D. Srinivasan, S. Mittal, P. Luethy and L. Kroos, unpublished data). . . . 70 . . . . . . . In addition to the ECF srgmas, the 0' family contains mne addrtronal putative Sigma factors, of which seven have been characterized, while the remaining two exhibit limited similarity to known sigma factors (Goldman et al. 2006, Kroos and Inouye 2008). The primary sigma factor, SigA, is essential for growth (Inouye 1990). SigB is a development-specific sigma factor that is essential for maturation of spores (Apelian and Inouye 1990). sigC is expressed early in development and affects fruiting body formation (Apelian and Inouye 1993). SigD is similar to the stationary-phase sigma factor, GS, of E. coli. M. xanthus sigD is present during vegetative growth and early in development. A sigD mutation results in growth defects, compromised stress responses, and abnormal development (Ueki and Inouye 1998). A sigD mutant is impaired in the synthesis of the intercellular A- and C-signals, but responds to starvation by inducing the synthesis of the stringent response molecule, (p)ppGpp (Viswanathan et al. 2006b). Si gE is present during vegetative growth and development, and is similar to heat shock sigma factors in other bacteria, but it does not appear to have a role in the heat shock response in M. xanthus (Ueki and Inouye 2001). SigF plays a role in S-motility and development (Ueki et al. 2005). SigG is similar to the E. coli FliA protein, which directs fiagellar gene expression. An M. xanthus SigG mutant displays no obvious defects in growth or development (Kroos and Inouye 2008). M. xanthus also has one 0'54 (Goldman et al. 2006), which is essential for growth, unlike in most bacteria that have been studied (Keseler and Kaiser 1997). Enhancer-binding proteins . . . 54 . . . . In bacterra, transcription at 0' promoters involves activators that bind to enhancer-like , 4 sequences upstream of the core promoter, and by DNA looping interact With 05 RNA polymerase (RNAP) to assist in open complex formation (Buck et al. 2000). These activators have a conserved central ATPase domain and a C-terminal DNA-binding domain. M. xanthus is predicted to have 52 genes that code for activators, based on the presence of the conserved ATPase domain (Goldman et al. 2006). These transcriptional activators have been designated (SM-activators (Gorski and Kaiser 1998), NtrC-like activators (Caberoy et al. 2003), or enhancer-binding proteins (EBPs) (Jakobsen et a]. 2004). In M. xanthus, most EBPs have an N-terminal sensory domain, suggesting a role in signal transduction. Gene knock-out and expression studies have demonstrated that 8' 1h 31 its certain EBPs play a role in aggregation and sporulation (Kroos and Inouye 2008). Some EBPs like N1a18 affect vegetative growth as well as development (Diodati et al. 2006). Fruiting body formation Nutrient depletion triggers the multicellular developmental process of M. xanthus. Deprivation of amino acids, carbon source, or phosphate initiates development (Manoil and Kaiser 1980b, Shimkets 1987, Dworkin 1996). Starvation conditions induce a rise in the intracellular concentration of the stringent response molecule (p)ppGpp (Manoil and Kaiser 1980a, Manoil and Kaiser 1980b Singer and Kaiser 1995). (p)ppGpp is essential for the initiation of the developmental process (Singer and Kaiser 1995). (p)ppGpp levels are affected by several genes and are subject to complex regulation. When starved at a high density, M. xanthus cells glide to form aggregation foci. Prior to and during the formation of aggregation centers, cells exhibit rippling behavior (Shimkets and Kaiser 1982). During rippling, cells accumulate in parallel ridges that appear to travel as waves under time-lapse microscopy (Welch and Kaiser 2001). As starvation continues, aggregation foci fuse and grow larger, resulting in the formation of mound-shaped structures over a period of 24 h (Kroos 2007). There are approximately 100,000 cells in a nascent fruiting body, and around 10-15% of the cells differentiate into heat and desiccation-resistant spherical spores, while other cells undergo programmed cell death (PCD) mediated by a toxin (Wireman and Dworkin 1977, Nariya and Inouye 2008). Cells that remain outside of the fruiting bodies are called peripheral rods (O'Connor and Zusman 1991). Peripheral rods are distinct from vegetative cells and they neither lyse nor sporulate. Their role in the developmental process remains to be understood. Intercellular signaling during development The process of development is controlled by temporal and spatial gene expression that is regulated by a network of intercellular signals and transcription factors. The involvement of intercellular signals in fruiting body development was discovered by the isolation of nonautonomous mutants that exhibit developmental defects (Hagen et al. 1978). The sporulation defects displayed by these mutants could be rescued by mixing with wild- type cells and this was defined as extracellular complementation (Hagen et al. 1978). The nonautonomous mutants were classified into five groups based on extracellular complementation experiments: asg (A-signal), bsg (B-signal), csg (C-signal), dsg (D- signal) and esg (E-signal) mutants (Hagen et a1. 1978, Downard et al. 1993). In these experiments, codevelopment of mutants from two different groups resulted in the rescue of developmental defects, while codevelopment of mutants from the same group did not show rescue (Hagen et a1. 1978, Downard et al. 1993). The signals are required at different times into the developmental process, with the A- and B-signals acting early in development at around 2 h, the D- and E-signals acting at 3-5 h, and the C-signal acting after 6 h (Kaiser 2004). The A- and C-signals have been studied in the most detail biochemically, while B-, D-, and E-signal molecules remain to be identified. A-signaling The A-signal is a mixture of extracellular proteases, peptides, and amino acids that is involved in sensing cell density early during the developmental process (Kuspa et al. 1992a, Kuspa et al. 1992b). Mutants that are defective in A-signal production have a defect in any one of the five asg genes (Kuspa and Kaiser 1989, Cho and Zusman 1999, Garza et al. 2000b). asgA encodes a hybrid protein containing an N-terminal receiver domain (typically found in RRs) and a C-terminal histidine kinase domain (Plamann et al. 1995). agsA is expressed during vegetative growth and during development. AsgA has autokinase activity and has been proposed to be involved in a phosphorelay process (Plamann et al. 1995). asgB codes for a putative DNA-binding protein that is essential for growth and development (Plamann et al. 1994). AsgB has a C-terminal helix-tum-helix domain that is similar to region 4 of the major sigma factors in M. xanthus, E. coli, and B. subtilis (Plamann et al. 1994). asgC encodes the major sigma factor of M. xanthus, sigA (Davis et al. 1995). asgD codes for a two-component signal transduction hybrid protein (Cho and Zusman 1999) similar to that encoded by asgA. ang encodes a protein with two putative membrane-spanning domains and shares homology with aminohydrolases (Garza et al. 2000a). The A-signal is comprised of heat-labile and heat-stable components (Kuspa et al. 19923, Plamann et al. 1992). The heat-labile component was shown to have trypsin-like protease activity. Inactivation of this activity by heat treatment resulted in 40 to 60% reduced activity of an early developmental gene at the 04521 locus (Plamann et al. 1992). It was Proposed that the heat-labile component of A-signal consists of secreted proteases required for degradation of proteins and peptides early in development. The heat-stable component was shown to primarily consist of amino acids; in particular, tyrosine, proline, phenylalanine, tryptophan, leucine, and isoleucine (Kuspa et al. 1992a). These and subsequent studies suggested that at the onset of starvation, cells release a mixture of proteases that act on outer membrane proteins to generate peptides and amino acids, and as starvation continues, increased levels of extracellular proteases, peptides, and amino acids result in induction of early developmental gene expression [reviewed in (Shimkets 1999, Kaiser 2004)]. The (24521 promoter was shown to be A-signal—dependent (Kuspa et al. 1986). This promoter is similar to 654-regulated promoters (Keseler and Kaiser 1995) and appears to be regulated by SasS and SasR, which are similar to HPKs and RRs respectively (Yang and Kaplan 1997). B-signaling A single locus, bsgA, is associated with all B-signaling mutants (Gill and Bornemann 1 988, Gill et al. 1988, Tojo et al. 1993b). The bsgA gene codes for an ATP-dependent intracellular protease which is very similar to Lon protease of E. coli (Gill et al. 1993, Tojo et al. 1993b, Tojo et al. 1993a). M. xanthus has two Lon proteases; LonV is essential for growth, and BsgA is essential for development (Tojo et al. 1993b). The mechanism of extracellular complementation is not known Because the BsgA protein is C ytoplasmic. 10 Interestingly, two suppressor mutations have been identified that bypass BsgA protease function during development. One mutation has been mapped to the sde gene that codes for an EBP (Hager et al. 2001) and another mutation has been mapped to the bcsA locus that codes for a protein similar to flavin-containg monooxygenases (Cusick et al. 2002). It appears that sde and bcsA might not function in the same pathway. .9de mutations also bypass the requirement for A-signaling but not that for C-signaling (Hager et a1. 2001), while mutations in bcsA suppress defects of C-signaling mutants but not of A- signaling mutants (Cusick et al. 2002). Recent work has shed some light on the possible function of the BsgA protease during development. It has been proposed to be involved in the cleavage of the transcriptional activator MrpC to an N-terminally truncated form, MrpC2, early in development (Nariya and Inouye 2006). MrpC2 appears to be a direct activator of fruA, a gene essential for fruiting body development (Ogawa et al. 1996, Ellehauge et al. 1998, Ueki and Inouye 2003). MrpC2 is undetectable in a bsgA mutant and fruA expression is also drastically reduced (Nariya and Inouye 2006). However, it remains to be tested whether MrpC is a direct substrate of the BsgA protease. Conceivably, MrpC2 and/or FruA activate transcription of one or more genes involved in production of the unknown B-signal. Alternatively, BsgA may cleave a protein other than MrpC that is involved in B- S i gnalin g. 11 D-signaling Similar to B-signaling, little is known about D-signaling, but a single locus, dsgA, has been implicated by genetic studies. dsgA codes for a putative translation initiation factor 3, IF3 (Cheng and Kaiser 1989b, Cheng and Kaiser 1989a, Cheng et al. 1994, Kalman et al. 1994). IF3 helps ribosomes in selection of initiation codons. M. xanthus dsgA complemented an E. coli infC (the gene for IF3) mutant (Cheng et al. 1994). DsgA is essential for M. xanthus growth and development (Cheng and Kaiser 1989b, Cheng and Kaiser 1989a). There was limited rescue of sporulation of dsgA mutants by wild-type cells upon co-development (Cheng and Kaiser 1989a), and the mechanism of extracellular complementation is unknown. E-signaling Mutations leading to defects in E-signaling have been mapped to two genes that code for the E101 and E113 subunits of branched-chain keto acid dehydrogenase (BCKDH) (Downard et al. 1993, Toal et al. 1995). esg mutants had aggregation and sporulation defects (Toal et a1. 1995). BKCDH is involved in conversion of branched-chain amino acids like leucine, isoleucine, and valine to isovaleryl-CoA, 2-methylbutyryl-COA, and isobutyryl-CoA, respectively, which are required for the synthesis of iso-fatty acids and Secondary metabolites (Ring et al. 2006). esg mutants have reduced iso-fatty acids and their developmental defects can be rescued by addition of isovalerate, suggesting that an iSo-fatty acid or a corresponding lipid could be required during development (Toal et al. 1 995). Kearns et al. showed the likely involvement of a phosphatidylethanolamine (PE) COntaining a straight-chain fatty acid (16:1(05c) in E-signaling (Kearns et al. 2001), 12 however, a recent study suggests that this PE is not required for M. xanthus development (Bode et al. 2006). Interestingly, the loss of BKCDH activity in esg mutants does not result in complete loss of iso-fatty acid production due to the presence of an alternative unique pathway in myxobacteria (Mahmud et al. 2002). A double mutant for both pathways is severely hampered in iso-fatty acid production (Bode et al. 2006). Developmental defects in this double mutant have been proposed to be due to the loss of unusual iso-branched ether lipids found in myxospores and therefore suggested to be novel biochemical markers of development (Ring et al. 2006). Whether extracellular complementation of esg mutants reflects developmental signaling or metabolite cross-feeding remains an open question, but the study of esg mutants has revealed an important role of iso-fatty acid biosynthesis in M. xanthus development. C-signaling C-signaling is mediated by the product of the csgA gene (Shimkets et al. 1983) and begins about 6 h after starvation (Kroos and Kaiser 1987). Mutants defective in C-signal synthesis are unable to ripple, aggregate, or sporulate (Shimkets et al. 1983). Developmental defects of csgA mutants can be rescued by either codevelopment with Wild-type cells or by exogenous addition of purified C-signal (Kim and Kaiser 1990a, I{im and Kaiser 1991). Li et a1. manipulated csgA expression in vivo to demonstrate that low levels of csgA expression induce rippling, intermediate levels induce aggregation, and high levels induce sporulation (Li et al. 1992). Kruse et al. showed that 13 In 313‘ [CI Sli (f) (”3 overexpression of csgA in vivo resulted in early aggregation, gene expression and sporulation (Kruse et al. 2001). These studies suggest that C-signaling coordinates the rippling, aggregation and sporulation behavior of cells. Initially, the C-signal was identified as a 17 kDa (p17) protein and purified by detergent extraction and biochemical fractionation of starving M. xanthus cells (Kim and Kaiser 1990d). The purified C-signal was able to restore development of csgA mutant cells (Kim and Kaiser 1990d). However, later findings suggested that the csgA gene product could possibly function as a short-chain alcohol dehydrogenase (SCAD) to produce the C- signal. The csgA gene encodes a 25 kDa (p25) protein that shares homology with SCADs and contains two conserved motifs, an N-terminal NAD+-binding pocket and a C- terminal catalytic domain, found in SCADs (Lee et al. 1995). csgA alleles carrying substitutions in either the putative N AD -brndrng pocket or catalytic domain were unable to complement csgA mutant cells (Lee et al. 1995). Likewise, exogenous addition of p25, purified after expression in E. coli, could rescue a csgA mutant, but there was no rescue with p25 having substitutions in either NAD+-binding pocket or catalytic domain (Lee et al. 1995). Also, overproduction of the SocA protein, which shares homology with SCADs, restores development of csgA mutant cells (Lee and Shimkets 1994). Recently, it was shown that p25 and p17 are two forms of the ngA protein. p25 is proteolytically cleaved to p17 by a serine protease in the starving cells and p17 lacks the DUtative NAD+-binding pocket (Lobedanz and Sogaard-Andersen 2003). p25 is present 14 Th. during vegetative growth, while p17 is present only during development (Kruse et al. 2001). Both p25 and p17 are cell-surface associated proteins (Lobedanz and Sogaard- Andersen 2003). p17 has been proposed to be perceived by an unidentified receptor on the adjacent cell (Figure 1.1). csgA expression is regulated in response to nutrient depletion. Li et al. showed that 930 bp upstream from the transcriptional start site (TSS) are required for full csgA expression in the presence of limited nutrients, while in the absence of nutrients 400 bp upstream of the TSS is sufficient for development and full csgA expression (Li et al. 1992). The products of the act operon have been shown to affect the level and timing of csgA expression (Gronewold and Kaiser 2001), but exactly how the Act proteins sense and respond to starvation is unknown. C-signal transduction requires cell alignment and possibly end-to-end contact between cells, and acts over a short range, communicating positional information (Kim and Kaiser 1990c, Kim and Kaiser 1990b, Kim and Kaiser 1990a, Sager and Kaiser 1994). As mentioned above, distinct levels of C-signaling are required for rippling, aggregation, and Sporulation. The expression of various genes during development also requires different levels of C-signaling (Kim and Kaiser 1991). The C-signal Transduction Pathway FruA plays an important role in the C-signal transduction pathway (Figure 1.1) and is believed to be regulated post-translationally by C-signaling (Ogawa et al. 1996, 15 Ellehauge et al. 1998). FruA regulates aggregation by activating Frz proteins (directly or indirectly) and FruA regulates sporulation by controlling expressron of at least 50 genes (Horiuchi et al. 2002). Starvation (p)ppGpp A-signal \ r it \ Aggr ation cg STPKs MrpC \|M and r—-D>——> HPK2 Sporulation T V\Mp6P—-IMEC:A Survival 74"“ _r:_ i\ 4:— p25 cs A mazF El—‘J FruA-P FruA-P 12 p17; Ser protease /HPK1 I" -*-<<]—i FruA < —- fruA Adjacent M. xanthus cells csgA Figure 1.1. Regulatory network during M. xanthus development. Posrtrve and negatrve effects (direct or indirect) are depicted by arrows and lines with barred ends, respectively. The C-signal (p17) is perceived by an unknown receptor on the surface of an adjacent cell and results in activation of an unknown kinase, HPK2. FruA has been proposed to be phosphorylated to a low level by an unknown HPK], independent of C Signaling, and later phosphorylated to a higher level by HPK2 in response to C srgnalrng HPK histidine protein kinase; STPKs, serine/threonine protein kinases; MrpC-P, phosphorylated MrpC; FruA-P, phosphorylated FruA. See text for detailed description fr‘uA transcription is induced around 6 h after the onset of starvation and depends on A- and E—signaling but not C-signaling (Ogawa et al. 1996, Ellehauge et al 1998) FruA IS 16 106 111; is fr it I"! similar to response regulators of the FixJ subfamily of two-component His-Asp signal transduction systems. The D59 residue in the N -terminal part has been proposed to be the site of phosphorylation and genetic evidence shows that it is indispensable for the function of FruA (Ellehauge et al. 1998). M. xanthus cells carrying D59A or D59N or D59Q mutant alleles were unable to complement aggregation and sporulation defects of a fruA mutant (Ellehauge et al. 1998). However, a D59E substitution designed to mimic the phosphorylated state of the protein was able to restore development of a fruA mutant (Ellehauge et al. 1998). A histidine kinase(s) that can phosphorylate FruA remains to be identified. The C-terminal domain of FruA consists of a helix-tum-helix DNA-binding motif (Ellehauge et al. 1998). The C-terminal DNA-binding domain of FruA (FruA- DBD) has been shown to bind to promoter regions of several developmental genes, including both C-signal-dependent and C-signal-independent genes. It has been proposed that early during development, FruA is phosphorylated to a small extent to activate transcription of C-signal-independent genes like dofA and tps, while later in development, as C-signaling increases, more FruA is phosphorylated to activate C-signal-dependent genes like fdgA, fmgA and dev (Ueki and Inouye 2005b, Ueki and Inouye 2005a, Yoder- Himes and Kroos 2006, Viswanathan et al. 2007b). fr‘uA expression is developmentally-regulated (Figure 1.1) and appears to be under the direct control of the development-specific transcription factor MrpC2 (Ogawa et al. 1996, El lehauge et al. 1998, Ueki and Inouye 2003). MrpC2 is an N-terminally truncated form 0f MrpC (Nariya and Inouye 2006). The mrpC locus was identified by transposon insertion mutagenesis as essential for aggregation and sporulation (Sun and Shi 2001b). l7 fps [1 11 11110 K \ prot: Mm! “th Mrp: 31.131 PR.) My PhOS fifth ' K. I Upstream of mrpC, there is a two—gene operon, mrpAB, that code for an HPK and a R. It has been shown that MrpB is required for mmC transcription and that MrpC autoregulates its own transcription (Sun and Shi 2001b). MrpC is similar to cyclic-AMP receptor family transcriptional activators. MrpC is present during vegetative growth, whereas MrpC2 is detectable only during development beginning at around 6 h (Nariya and Inouye 2006). BsgA, an ATP-dependent protease has been proposed to cleave off the 25 N-terminal residues of MrpC to produce MrpC2 (Figure 1.1). A serine-threonine protein kinase (STPK) cascade controls the activity of MrpC and the generation of MrpC2. Pkn8, an integral membrane STPK, phosphorylates a cytoplasmic STPK, Pknl4, which then phosphorylates MrpC (Nariya and Inouye 2005). Phosphorylation inhibits processing of MrpC to MrpC2. MrpC2 cannot be phosphorylated, suggesting that Thr21 and/or Thr22 are the likely sites of MrpC phosphorylation. Phosphorylation reduces MrpC DNA—binding activity, and MrpC2 binds to the mrpC and fruA promoter regions with higher affinity than does MrpC (Nariya and Inouye 2006). The inhibitory effects of the Pkn8-Pkn14 cascade are relieved by an unknown mechanism early during development, causing a rise in the levels of MrpC and eventually MrpC2. MrpC plays an important role in cell fate decisions during development (Figure 1.1). MrpC was shown to act as an anti-toxin that interacts directly with the toxin, MazF, an mRNA interferase, which mediates PCD during development (Nariya and Inouye 2008). MrpC also binds to the mazF promoter region and appears to activate transcription. I)hOSphorylation of MrpC affects its antitoxin properties as it has been shown that 1“onphosphorylated MrpC has higher affinity for MazF compared to the phosphorylated 18 A form SIP] wit form of MrpC (N ariya and Inouye 2008). During growth, MrpC is phosphorylated by the STPK cascade, but MazF is kept in check Because phosphorylated MrpC presumably activates mazF transcription poorly. During development, MrpC and/or MrpC2 presumably activate mazF gene expression, leading to PCD of some cells in the population. MrpC and/or MrpC2 presumably inhibit MazF in other cells, preventing PCD and activating genes required for aggregation and sporulation. Binding of MrpC2 to the mazF promoter region and MazF has not been tested, but it is important to do so Because processing of MrpC to MrpC2 could be an important determinant of whether a cell undergoes PCD or forms a spore. C—signal-dependent gene expression De velopmentally-regulated promoters in M. xanthus have been identified by transposition of a T n5 lac into M. xanthus chromosome, generating transcriptional fusions of the promoter-less E. coli lacZ gene to genomic regulatory regions (Kroos and Kaiser 1984). Each insertion locus was assigned a four digit number. These fusions proved extremely useful for studying developmental gene expression in response to intercellular signals. Of the 2,374 insertions, 29 fusions were found to be developmentally-regulated (Kroos et a1. 1 986). In one study, 18 fusions were found to be A-signal-dependent (Kuspa et al. 1986). In another study, 26 insertions were found to be B—signal-dependent and 15 were C- Si gnal-dependent (Kroos and Kaiser 1987). The Kroos lab has cloned DNA upstream of SeVeral C-signal-dependent Tn5 lac insertions, mapped transcriptional start sites, I>erf()I‘med deletion analysis to identify promoter regions and conducted extensive mutational analysis to identify cis-regulatory sequences (Fisseha et al. 1996, Brandner 19 and Kroos 1998, Fisseha et al. 1999, Viswanathan and Kroos 2003, Yoder and Kroos 2004a, Yoder and Kroos 2004b, Loconto et al. 2005, Viswanathan et al. 2006a). Sequence comparison and mutational analysis identified two conserved Cir-acting sequences, the C box (consensus CAYYCCY; Y is C or T) and the 5-bp element (consensus GAACA) (Fisseha et al. 1999, Viswanathan et al. 2006a). These sequences are present in all C-signal-dependent promoter regions studied so far and are crucial for promoter activity (Fisseha et al. 1996, Brandner and Kroos 1998, Fisseha et al. 1999, Viswanathan and Kroos 2003, Yoder and Kroos 2004a, Yoder and Kroos 2004b, Loconto et al. 2005, Viswanathan et al. 2006a). These studies have provided a foundation for understanding the regulation of C-signal-dependent gene expression. Mutational analysis of the (24403 promoter region revealed that a 10-bp element, in addition to the C box and the 5-bp element, is critical for expression (Viswanathan and Kroos 2003). The 04403 fusion is absolutely dependent on C-signaling Because expression is completely abolished in a csgA mutant background but is restored upon codevelopment with wild-type cells, which supply C-signal (Kroos and Kaiser 1987). The 04400 promoter region has a 5-bp element and a C box at exactly the same position as present in the (24403 promoter region (Brandner and Kroos 1998, Yoder and Kroos 2004a). However, single base-pair mutations in these sequences have different effects on Promoter activity (Viswanathan and Kroos 2003, Yoder and Kroos 20043). A region from ~101 to +155 is sufficient for 04400 promoter activity (Yoder and Kroos 2004a). FruA is indispensable for expression from the (24400 promoter region (Yoder-Himes and Kroos 20 2006). FruA-DBD binds to a region between -86 and -77, which contains a region (-86 to -81) that is responsible for partial dependence on C-signaling (Yoder-Himes and Kroos 2006). 0'A RNA polymerase, the major form of vegetative RNA polymerase, was unable to produce transcripts from the (24400 promoter (Brandner and Kroos 1998). Null mutations in SigB and sigC genes did not affect expression from the (24400 promoter (Brandner and Kroos 1998). However, mutations in sigD and sigE abolished and reduced Q4400 promoter activity, respectively, suggesting a direct or indirect role of SigD and SigE in 04400 transcription (Yoder and Kroos 2004a). There are two C boxes and two 5-bp elements present in the 04499 promoter region, which are crucial for the Q4499 promoter activity (Yoder and Kroos 2004b). Single base- pair mutations in one of the 5-bp elements and one of the C boxes had different effects on promoter activity than mutatiOns at corresponding positions in the (24400 and (24403 promoter regions, suggesting a difference in the interaction with trans-acting factors. A region from -100 to +50 contains the regulatory sequences required for 04499 promoter activity (Yoder and Kroos 2004b). Expression from a 04499 fusion, like that from a 04400 fusion, is partially dependent on C-signaling (Kroos and Kaiser 1987). Also, like the 04400 promoter, O'A RNA polymerase was unable to produce transcripts from the 04499 promoter (Fisseha et al. 1999). There was no effect of sigB or sigC null mutations on expression from the 04499 promoter (Brandner and Kroos 1998). A null mutation in SigD and sigE reduced expression from the 04499 promoter to about 30% of the level Observed in wild-type cells (Yoder and Kroos 2004b). This suggests that sigD and sigE directly or indirectly affect Q4499 promoter activity. 21 Erpr Inter elem Cho Expression of lacZ from the 04406 Tn5 lac fusion depends absolutely on C-signaling (Kroos and Kaiser 1987, Loconto et a1. 2005). An upstream negative regulatory element mediates C-signal dependence of the 04406 promoter (Viswanathan et al. 2006a). Interestingly, DNA 50 to 140 bp downstream of the TSS contains a positive regulatory element. Similar to other C—signal-dependent promoter regions, the 5-bp element and the C box are important for 04406 promoter activity (Viswanathan et al. 2006a). The dev operon, whose products are essential for sporulation, is partially dependent on C- signaling (Kroos et al. 1986, Kroos and Kaiser 1987, Kroos et al. 1990). The regulatory region controlling dev expression spans more than 1 kb and includes positive and negative regulatory elements surrounding the promoter (Viswanathan et al. 2007a). The dev promoter region has the 5-bp element and C box-like sequences, but mutational analysis has not been performed on these sequences. dev expression is under combinatorial control by FruA and LadA (Viswanathan et al. 2007b). FruA binds to an upstream positive regulatory region centered at -91, while LadA binds around 350 bp downstream of the dev promoter (Viswanathan et al. 2007b). A 5-bp element and a C box are present in the fruA promoter region, although fruA transcription is C-signaling-independent (Ellehauge et al. 1998, Srinivasan and Kroos 2004). Mutational analysis has shown that these sequences are important for fruA CXpression (Srinivasan and Kroos 2004). 22 id: id: fr”. To understand differential gene expression in response to C-signaling at the molecular level, my research focused on elucidating the regulation of two developmentally- regulated C-signal-dependent transcription units, fingA (EruA- and MrpC2-regulated gene _A_) and fmgBC (EruA- and MrpC2-regulated gene B and Q). fingA and fingBC genes were identified by insertion of Tn5 lac into the M. xanthus genome at loci 04400 and (24499, respectively (Kroos et al. 1986). Chapter 2 describes the identification of MrpC2 as a direct activator of fingA expression. It was found that fingA expression is under combinatorial control of the previously identified activator, FruA, and MrpC2. FruA is essential for MrpC and/or MrpC2 association with the fmgA promoter region in vivo. This is a novel mechanism of gene regulation in which a response regulator, FruA, recruits an independent transcription factor, MrpC2. Biochemical studies showed that FruA and MrpC2 bind cooperatively to the fmgA promoter region. We propose that cooperative binding subjects fingA promoter activity to positional information via C-signaling and phosphorylation of FruA, and to starvation signaling via MrpC2 during development. This chapter will be submitted to Proceedings of National Academy of Sciences in August 2008. In Chapter 3, regulation of fingBC operon expression is described. MrpC2 and FruA were identified as direct activators of fingBC expression since both proteins bind to important Cis-regulatory sequences in the fingBC promoter region. Similar to fmgA, cooperative binding of FruA and MrpC2 was observed at the fingBC promoter region. This chapter Will be submitted to the Journal of Bacteriology in September 2008. 23 Chapter 4 is conclusions and future directions, which contains a summary of the research and possible future directions. 24 Chapter 2: Cooperative binding of a response regulator and a bifunctional transcription factor/antitoxin controls gene expression and cell fate during Myxococcus xanthus development The work in this chapter will be submitted to Proceedings of National Academy of Sciences in August 2008. 25 ABSTRACT Myxococcus xanthus is a bacterium that undergoes multicellular development requiring coordinate regulation of multiple signaling pathways. One pathway governs aggregation and sporulation of some cells in a starving population and requires C—signaling, while another pathway causes programmed cell death and requires the MazF toxin. In response to starvation, the levels of the antitoxin MrpC and its related proteolytic fragment MrpC2, are increased, inhibiting the cell death pathway via direct interaction of MrpC with MazF. Herein, we demonstrate that MrpC2 plays a direct role in the transcriptional response to C-signaling. We show that MrpC2 binds to sequences upstream of the C-signal- dependent fingA promoter. These sequences are present in other C-signal-dependent promoter regions, indicating a general role for MrpC2 in developmental gene regulation. Recruitment of MrpC2 to promoters is enhanced by FruA, a protein that responds to C- signaling and is similar to response regulators of two-component signal transduction systems. DNA binding studies showed that this involves a novel mechanism for a response regulator, in which FruA and MrpC2 bind cooperatively to adjacent sites upstream. of the fmgA promoter. This novel mechanism of combinatorial control allows coordination of morphogenetic C-signaling via FruA with starvation signaling and cell death via MrpC2 and MrpC, determining spatiotemporal gene expression and cell fate. 26 INTRODUCTION Understanding how cells integrate many different signals to regulate genes and determine cell fates during multicellular development is a fundamental question. Myxococcus xanthus provides an attractive model to address this question because starvation initiates a relatively simple developmental process (Whitworth 2008). Thousands of rod-shaped cells coordinate their movements to build fruiting bodies in which cells differentiate into dormant, spherical spores (Figure 2.1). However, not all cells form spores. Alternative fates are programmed cell death (PCD) (Nariya and Inouye 2008) or persistence outside of fruiting bodies as peripheral rods (O'Connor and Zusman 1991). Signals act at different times during the M. xanthus developmental process to control gene expression, coordinate cell movements, and determine cell fates (Kroos 2007). Starvation triggers the stringent response, which involves production of the second messenger (p)ppGpp (Figure 2.1). This intracellular signal leads to activation of early developmental genes (like csgA) (Harris et al. 1998, Crawford and Shimkets 2000) and secretion of protease activity that generates a mixture of peptides and amino acids known as A-signal (Kuspa et al. 1992a, Plamann et al. 1992). This extracellular signal is believed to allow quorum sensing (Kuspa et al. 1992b). At a sufficient concentration, expression of genes like those in the mrp operon is induced (Figure 2.1) (Sun and Shi 2001a). Later, when cells begin to aggregate, C-signaling takes over. The C-signal appears to be a proteolytic cleavage product of ngA, which is associated with the cell surface (Kim and Kaiser 1990a, Lobedanz and Sogaard-Andersen 2003), but a receptor has not been identified. C-signaling requires cell alignment (Kim and Kaiser 1990b) and 27 possibly end-to-end contact (Sager and Kaiser 1994), so it is paracrine or short-range signaling, which is common in eukaryotes but rare among bacteria (Bassler and Losick 2006). The short-range nature of C-signaling and its effects on cell movement and gene expression can explain its critical role in coordinating aggregation with sporulation (Kaiser 2003, Sogaard-Andersen et al. 2003). Cell alignment within a nascent fruiting body has been proposed to allow a high level of C-signaling and activation of genes required for sporulation (Sager and Kaiser 1993). Consistent with this model, elevated ngA accelerates aggregation and sporulation, but the fruiting bodies are smaller than normal, and reduced ngA causes the opposite effects (Li et al. 1992, Gronewold and Kaiser 2001, Kruse et al. 2001). Also, the expression of genes important for sporulation is temporally and spatially restricted to nascent fruiting bodies (Sager and Kaiser 1993, Julien et al. 2000). Here, we focused on gene regulation in response to C-signaling during M. xanthus development, a process in which FruA plays a key role (Figure 2.1). FruA is similar to response regulators of two-component signal transduction systems (Ogawa et al. 1996). FruA’s N-terminal domain is believed to be phosphorylated by one or more histidine protein kinases (HPKs) in response to C-signal and perhaps other signals (Ellehauge et al. 1998, Jelsbak et al. 2005), but the cognate HPK(s) remains to be identified. Phosphorylation presumably enhances DNA binding by the C-terminal domain of FruA, which resembles that of the NarL/FixJ subfamily of response regulators (West and Stock 2001). The FruA DNA-binding domain (FruA-DBD) has been shown to bind to sites in the promoter regions of developmental]y-regulated genes, including one whose 28 expression does not depend on C-signaling (dofA; Figure 2.1) (Ueki and Inouye 2005a) and others whose expression is C-signal-dependent (e.g., fmgA; Figure 2.1) (Ueki and Inouye 2005b, Yoder-Himes and Kroos 2006). FruA positively regulates expression of these genes, but in the case of fingA, mutational analysis of the promoter region implied that an additional transcriptional activator is required (Yoder and Kroos 2004a). The fingA gene (named herein) was identified by insertion of transposon Tn5 lac into the M. xanthus genome at locus $24400 (Kroos et al. 1986). Expression of lacZ from Tn5 lac 524400 was shown to be developmentally-regulated and partially dependent on csgA (Kroos et al. 1986, Kroos and Kaiser 1987). M. xanthus DNA upstream of Tn5 lac 94400 was cloned, a putative transcriptional start site was mapped, and the region from - 101 to +155 was shown to encompass the fingA promoter (Brandner and Kroos 1998). Expression from this promoter region, as measured by a transcriptional fusion to lacZ integrated ectopically into the M. xanthus chromosome, was reduced in a csgA mutant (as was expression from T n5 lac £24400), but expression was restored upon co-development of the csgA mutant with wild type cells, which supply C-signal. This indicated that fingA promoter activity is partially dependent on C-signaling. Mutational analysis identified cis-regulatory sequences at -86 to -77 and -63 to -46 upstream of the fingA promoter (Figure 2.2) (Yoder and Kroos 2004a), and subsequent analysis showed that FruA-DBD binds to the sequence between -86 and -77 (Yoder-Himes and Kroos 2006). The sequence between -63 and -46 contains two elements found in other C-signal-dependent promoter regions, a 5-bp element (consensus GAACA) and a C box (consensus CAYYCCY; Y means C or T) (Fisseha et al. 1999, Viswanathan and Kroos 2003, Yoder and Kroos 29 300 iii ’19-1 wit. Tilft' 2004a, Loconto et al. 2005, Viswanathan et al. 2006a, Viswanathan et al. 2007a). Mutations in this region nearly abolish fingA promoter activity (Figure 2.2) (Yoder and Kroos 2004a) but do not affect binding of FruA-DBD (Yoder-Himes and Kroos 2006). Taken together, these studies suggested that a transcriptional activator binds to the sequence between -63 and -46 upstream of the fingA promoter, and perhaps to similar sequences in other C-signal-dependent promoter regions. Here, we report identification of the activator as MrpC2 and we name the gene at the 524400 locus fingA (EruA- and MrpC2-regulated gene A). MrpC2 lacks the N-terminal 25 residues of MrpC and appears to be generated by proteolytic activity of a developmental]y-regulated protease (LonD; Figure 2.1) (Nariya and Inouye 2006). MrpC is similar to transcription factors in the cyclic AMP receptor protein (CRP) family (Sun and Shi 2001b). Recently, MrpC was shown to interact with the toxin MazF, which mediates PCD during development (Nariya and Inouye 2008). Phosphorylation of MrpC by a cascade of serine/threonine protein kinases (STPKs) appears to inhibit its proteolytic processing to MrpC2 during growth (Nariya and Inouye 2005, Nariya and Inouye 2006). Early during development, due to relief from the inhibitory effects of the STPK cascade, MrpC and MrpC2 levels are increased. MrpC2 has a higher affinity than MrpC for binding sites in the mrpC and fruA promoter regions, so MrpC2 presumably boosts transcription from these promoters. In addition to identifying MrpC2 as an activator of fingA transcription, we show that FruA is required for association of MrpC and/or MrpC2 with the fingA promoter region in vivo, and that FruA and MrpC2 bind cooperatively to fmgA promoter region DNA in vitro. Cooperative binding of a response regulator with 30 another transcription factor represents a novel mechanism of gene regulation. Preliminary results indicate that this mechanism is shared by other C-signal-dependent genes (see Discussion). We propose that cooperative binding facilitates integration of positional information (via short-range C-signaling through FruA) with nutritional status and other signals (through transcriptional and post-transcriptional control of MrpC), governing cell fate decisions, analogous to combinatorial control during development of multicellular eukaryotes. 31 MATERIALS AND METHODS Bacterial strains and plasmids Strains and plasmids used in this study are listed in Table 2.1. Growth and development E. coli BL21(DE3) containing plasmids were grown at 370C in Luria-Bertani medium (Sambrook et al. 1989) containing 200 ug ampicillin per ml. Growth and development of M. xanthus was as described (Viswanathan and Kroos 2003). Preparation of fmgA DNA fragments DNA fragments spanning the fingA promoter region from -101 to +25 were generated by PCR using wild-type or mutant plasmid as the template and the primers 5’- C'ITAAGC'I'ITGCACTGCGACGCGAGTC-3' (for -101) and 5 '- GCGGATCCCGGTCC’ITCGCGTCGCCG-3' (for +25). For EMSAs, 32P-labeled DNA was synthesized by PCR after labeling the primers with [7'32P]ATP using T4 polynucleotide kinase, and the labeled DNA was purified and the concentration was estimated as described (Yoder-Himes and Kroos 2006). For DNase I footprinting, only one primer was labeled and a different upstream primer, 5'- AGGCT'I'I‘CGATGCACTGCG-3' (for -139), was used to allow resolution of digestion products of interest. The -139 to +25 DNA fragment produced a pattern of shifted complexes in EMSAs with Hile-MrpC2 and/or FruA-His6 that was indistinguishable from the -101 to +25 DNA fragment (data not shown). The modified (+5 bp) -76 to -41 32 fragment was generated by annealing 5'- MTGCGGTGGGGAGCGAACAGTCCCACATCCCTGGCGG-3' (the non-fmgA sequence is underlined) with its complement; after each oligonucleotide had been labeled as above, they were mixed, boiled for 10 min, placed at room temperature for 3 h, then the double-stranded DNA fragment was purified as above. EMSAs and DNase I footprinting EMSAs were performed as described (Yoder-Himes and Kroos 2006) except the binding reaction mixtures were incubated at 250C for 15 min. For footprinting, 0.2 U of DNase I (Promega) was added to the binding reaction mixture (20 [.ll) for 2 min at 250C. The binding mixture was as described previously (Yoder-Himes and Kroos 2006), except that it included SmM MgClz, 0.5 mM CaClz, 0.025 rig/[1.1 double-stranded poly(dI-dC), and no glycerol. Reactions were stopped by adding 100 11.1 of solution containing 300 mM sodium acetate, 20 mM EDTA, 0.2% SDS, 0.02 ug/ul proteinase K, and 100 ug/ml yeast tRNA, and incubating at 520C for 15 min. After extraction with 100 [.11 of phenol (twice), DNA was precipitated with ethanol. The DNA was resuspended in formamide loading buffer (Sambrook et al. 1989), boiled for 3 min, subjected to electrophoresis on an 8% polyacrylamide gel containing 8 M urea, and visualized by autoradiography. Sequencing ladders were generated using the SequiTherm EXCELTM H DNA Sequencing Kit protocol (Epicentre Biotechnologies). 33 DNA-affinity chromatography An fmgA DNA fragment (-101 to +25) was synthesized by PCR with a 5'-biotin label at - 101, bound to streptavidin beads, and DNA-affinity chromatography was performed with the AS fraction as described (Viswanathan et al. 2007b). Antibody supershift assays Binding reactions were performed as described above and then antibodies were added and the mixture was incubated at 40C for 30 min, followed by EMSAs as described (Yoder-Himes and Kroos 2006). Preparation of MrpC2 and FruA His lo-MrpCZ (Nariya and Inouye 2006) and FruA-DBD-Hiss (Yoder-Himes and Kroos 2006) were purified as described previously from E. coli strains SMhisMrpC2 and EDYFruA, respectively. FruA-Hi36 was purified from E. coli SMFruAhis as described (S. Inouye, personal communication). ChIP M. xanthus strains MDY4400.DZF 1 and MDY4400.FA were used for ChIP as described (Yoder-Himes and Kroos 2006) with the following modifications: anti-MrpC antibodies (500 ng) (Nariya and Inouye 2006) or control IgG (500 ng) (Santa Cruz Biotechnology) were used for immunoprecipitation, twofold serial dilutions were made of the input DNA samples, and the primers used for PCR of the fingA promoter region were the one for +25 34 described above and one upstream (yielding a product of about 180 bp) in the vector used for ectopic integration (5'-CTGCCAGGAATI‘GGGGATC-3’). 35 TABLE 2.1. Bacterial strains and plasmids used in this study Bacterial strain or plasmid E. coli strains BL21(DE3) SMhisMrpC2 SMFruAhis EDYFruA M. xanthus strains DKl622 DK4292 MDY4400.DZF 1 MDY4400.FA Plasmids pETl 1a/FDBD- H8 pETl6b/Hislo- MrpC2 pETl 1km/FruA- His6 pJB4001 pJB40029 pDY69 pD Y79 pD Y67 Description F‘ ompT hsdsBoB' mB') gal dcm with DE3, a it prophage carrying the T7 RNA polymerase gene BL21(DE3) containing pET16b/His lo-MrpCZ BL21(DE3) containing pETl 1km/FruA-His6 BL21(DE3) containing pETl lalFDBD-Hg Wild type Tn5 lac $24400 sglAI attB::pJB40030 sglAI fruA::TnV 0786 attB : :pJ B40030 pETl 1a with a gene encoding FruA- DBD-I-Ii33 under control of a T7 RNA polymerase promoter pETl6b with a gene encoding Hisw- MrpC2 under control of a T7 RNA polymerase promoter pETl 1km with a gene encoding FruA-11136 under control of a T7 RNA polymerase promoter pGEM7Zf with the 1.1 kb EcoRI- BamHI fragment from pJB4400 pGEM7Zf with fmgA DNA from -101 to +155 pJB40029 with GT C-to-TGA mutation at —86 to —84 pJB40029 with GGGGGTG-to- T'ITI'TGT mutation at --83 to —77 pJB40029 with TG-to-GT mutation at —76 to -75 36 Reference or Source Novagen This work This work (Yoder-Himes and Kroos 2006) (Kaiser 1979) (Kroos et al. 1986) (Yoder-Himes and Kroos 2006) (Yoder-Himes and Kroos 2006) (Ueki and Inouye 2005b) (Nariya and Inouye 2006) S. Inouye (Brandner and Kroos 1998) (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) pDY37 pDY65 pDY35 pDY61 pDY59 pDY57 pGV4400. 1 Table 2.1 (cont’d) pJ B40029 with CGGTG-to-A'ITGT mutation at -74 to —70 pJB40029 with GGGAGC-to- TITCTA mutation at —69 to —64 pJB40029 with GAAC-to-TCCA mutation at —63 to —60 pJB40029 with A-to-C mutation at — 59 pJB40029 with GTCCC-to-TGAAA mutation at -58 to —54 pJB40029 with A-to-C mutation at — 53 pJB40029 with CATCCCT-to- ACGAAAG mutation at -52 to —46 37 (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) (Yoder and Kroos 2004a) RESULTS An insertion in fmgA delays aggregation Tn5 lac 524400 in M. xanthus DK4292 is inserted in codon 138 of an open reading frame predicted previously to code for a protein that binds ATP or GTP (Brandner and Kroos 1998). The genomic sequence of M. xanthus revealed a long open reading frame (MXAN2884) preceded by a gene in the opposite orientation and followed closely by five genes in the same orientation that may form an operon (Goldman et al. 2006). The predicted start codon of MXAN2884 (Goldman et al. 2006) is likely incorrect, as it is not preceded by a good potential ribosome-binding site and it is located at -79 relative to the mRNA 5’ end mapped previously (Brandner and Kroos 1998), which mutational analysis confirmed as the fingA transcriptional start site (Yoder and Kroos 2004a). On the other hand, Brandner and Kroos (Brandner and Kroos 1998) identified a potential ATG start codon beginning at +66 that is preceded 5 bp upstream by the sequence AGGGAGG, which is a good potential ribosome-binding site since it is complementary (except for one mismatch) to a sequence near the 3’ end of M. xanthus 16S rRNA (Oyaizu and Woese 1985). The ATG beginning at +66 is likely the correct start codon of MXAN2884, which we name FmgA herein. A BLAST search with the predicted FmgA sequence revealed two putative domains; an N-terminal ATPase domain and a C-terminal tetratricopeptide repeat domain. The N-terminal domain was most similar to proteins found in other myxobacteria (a putative STPK of Sorangium cellulosum [e45] and a putative adenylyl 0!“ guanylyl cyclase of Stigmatella aurantiaca [e'lo] that also had a C-terminal tetratricopeptide repeat domain). 38 starvation / l (p)ppGpp “few @®\ 0 ./ @A- -signal -+ ll 1': WP |mrpA mrpB .® w]... . mazF I 1 ® @l—fl l [Tr—‘1 fmA l : C- signal—>. ®::.-Q—:D:::1 cell death .IIII CI... dofA V l-I__'] fmgA Figure 2.1. Model of the M. xanthus regulatory network. Effects depicted in the network may be direct or indirect. The mrpA and mrpB genes are co-transcribed from a promoter that depends on a cascade of enhancer-binding proteins (EBPs) (N. Caberoy & A.G. Garza, personal comm), which activate transcription by 0 RNA polymerase. MrpA is a putative histidine protein kinase (HPK) thought to phosphorylate MrpB, a putative EBP hypothesized to activate mrpC transcription (Sun and Shi 2001b). Phosphorylation of MrpC depends on a cascade of serine/threonine protein kinases (STPKs) (Nariya and Inouye 2005). Other effects are described in the text. 39 M. xanthus DK4292 bearing Tn5 lac £24400 inserted in fingA exhibited delayed aggregation by about 6-h compared to wild-type DKl622 but both strains formed a similar number of heat- and sonication-resistant spores that were able to germinate and form a colony (data not shown). The effect on aggregation could be due to loss of FmgA and/or due to loss of one or more downstream gene products (due to a polar effect of the Tn5 lac insertion) that might also be transcribed from the fingA promoter. MrpC2 binds to a cis-regulatory sequence in the fmgA promoter region As described above, mutational analysis identified a cis-regulatory sequence located at - 63 to -46 upstream of the fmgA promoter (Figure 2.2) (Yoder and Kroos 2004a). This sequence contains two elements found in other C-signal-dependent promoter regions, a 5- bp element and a C box, suggesting that these elements might be bound by a transcriptional activator of fingA that also regulates other C-signal-dependent genes. To identify the putative activator, DNA—binding proteins were partially purified as described previously (Ueki and Inouye 2003) from M. xanthus that had undergone 12 h of development, a time when fingA is expressed (Kroos et al. 1986). Proteins in the AS fraction were incubated with a 32P-labeled DNA fragment (-101 to +25) spanning the fingA promoter region and electrophoretic mobility shift assays (EMSAs) revealed a single shifted complex (Figure 2.2). EMSAs with DNA probes having a mutation between -63 and -46 eliminated or greatly reduced formation of the shifted complex, with the exception of the single base pair change at -53, which also had a smaller effect on promoter activity in vivo (Figure 2.2) (Yoder and Kroos 2004a). The shifted complex 40 appeared to be formed by a protein in the AS fraction that binds specifically to sequences between ~63 and -46 upstream of the fmgA promoter. FruA site 5-bp element C-box ~86 @bGGGGTdTGQGGTEbGGAGCmETCCaA ATCCC -46 l GT ATTGT TlggTA I 92 TI'T'TGT TGA 51 C v 26 TCCA C TGAAA 15 ACGAAAG i+.x Figure 2.2. Effects of mutations on fingA promoter activity in vivo and on DNA binding in vitro. The top part summarizes mutational effects on developmental fmgA-lacZ expression (Yoder and Kroos 2004a). The wild—type fingA upstream sequence is alternately underlined or boxed to indicate mutations. Mutant sequences are shown below. The number beneath each mutant sequence indicates the percentage of wild—type promoter activity. The bottom part shows EMSAs performed with P-labeled fmgA DNA (6 nM) spanning from -101 to +25 and proteins in the AS fraction (0.7 ug/ul). The arrow indicates the shifted complex produced by incubating the wild-type (WT) DNA fragment with the AS fraction, and other lanes show the effects of mutations. To purify the putative activator protein from the AS fraction, DNA-affinity chromatography was performed with the fmgA promoter region (-101 to +25). The major protein species after purification had an apparent molecular weight of about 30 kDa (Figure 2.3A). The affinity-purified protein (APP) generated a shifted complex 41 indistinguishable from that observed with the AS fraction (Figure 2.3B). Also, like the AS fraction (Figure 2.2), APP failed to generate a shifted complex with mutant (~63 to - 60) fingA promoter region DNA (Figure 2.3B). Therefore, APP was subjected to mass spectrometry analysis after protease digestion. Peptide sequences matching MrpC were the only significant matches to M. xanthus proteins predicted from the genome sequence. MrpC is similar to CRP-family transcription factors and was shown previously to be essential for development (Sun and Shi 2001b). A form of MrpC lacking the N-terminal 25 residues, called MrpC2, was identified previously in an AS fraction based on binding ' to the fruA promoter region (Ueki and Inouye 2003). Our results suggested that MrpC2 in the AS fraction binds to the fingA promoter region at a site (~63 to ~46) near the previously identified FruA binding site (~86 to ~77) (Yoder-Himes and Kroos 2006). To test the idea that MrpC2 in the AS fraction was responsible for the shifted complex (Figure 2.2), antibodies against MrpC were added after the complex had been allowed to form. EMSAs revealed the formation of supershifted complexes and loss of the original shifted complex (Figure 2.4), supporting the idea that MrpC2 in the AS fraction binds to fingA promoter region DNA, producing the original shifted complex. 42 ' —105 _78 WT mutant —55 APP ' ' ' + ' + —45 - —34 1 "17 _ _—16 Figure 2.3. DNA-affinity purification of protein that binds to the fingA promoter region. (A) SDS-PAGE of protein purified from the AS fraction using fmgA DNA (-101 to +25). The arrow indicates the major species in the affinity-purified protein (APP) after staining with silver. Numbers indicate the migration positions of molecular weight (kDa) standards. (B) EMSAs with 32P-labeled fingA DNA (6 nM) spanning from —101 to +25 and proteins in the AS fraction or the APP. The arrow indicates the shifted complex produced with the wild-type (WT) DNA fragment. APP failed to form the shifted complex with a DNA fragment bearing the GAAC to TCCA mutation at ~63 to ~60 (mutant). 43 G-Ml’pC A AS-++++ Pt; Figure 2.4. Anti—MrpC antibodies supershift the complex formed by fingA promoter region DNA and the AS fraction. 32P-labeled fmgA DNA (6 nM) spanning from ~101 to +25 was incubated with the AS fraction followed by addition of 0.03 uM, 0.06 11M, or 0.1 uM anti-MrpC antibodies. The shifted complex is indicated by the arrow and supershifted complexes are indicated by the bracket. To confirm that MrpC2 binds to the fmgA promoter region, N-terminally His-tagged MrpC2 (His lo—MrpCZ) was expressed in E. coli and purified. Hislo~MrpC2 exhibited a similar pattern of binding to wild-type and mutant fingA promoter region DNA (~101 to +25) as seen with the AS fraction (Figure 2.5). The complex produced by His 10—MrpC2 migrates more slowly than the complex produced by the AS fraction, presumably due to the 10 His residues plus 8 additional residues present in the His10~MrpC2 fusion protein. Mutations between ~63 and ~46 eliminated or reduced MrpC2 binding, with the exception of a single base pair change at ~53. These results, taken together with the effects of mutations in this region on fmgA promoter activity (Figure 2.2) (Yoder and Kroos 2004a), imply that MrpC2 binding to this region activates fmgA transcription. Since the region includes a 5-bp element and a C box, which are found in a similar arrangement, separated by 5-8 base pairs, upstream of other C-signal-dependent promoters (Fisseha et al. 1999, Viswanathan and Kroos 2003, Yoder and Kroos 2004b, Loconto et al. 2005, Viswanathan et al. 2006a, Viswanathan et al. 2007a), another implication is that MrpC2 might directly activate other C—signal-dependent genes (see Discussion). Interestingly, mutations upstream of ~63 appeared to enhance (~74 to ~70) or reduce (~76 to ~75) MrpC2 binding, whereas two mutations between ~86 and ~77 that impair binding of FruA-DBD (Yoder-Himes and Kroos 2006) did not affect MrpC2 binding (Figure 2.5). We conclude that FruA and MrpC2 bind to adjacent, important cis-regulatory sequences upstream of the fmgA promoter. The MrpC2 binding site in the fingA promoter region is the third type of MrpC2 binding site to be described. It includes a 5-bp element and a C box, but other sequences between ~76 and ~46 also affected binding (Figures 2.2 and 2.5), so MrpC2 interacts with a long segment of DNA at this type of site. Two other types of MrpC2 binding sites have been described previously (consensus sequences GTGTC-Ng-GACAC and G/A'I'I'I'C/GAA/G) (Nariya and Inouye 2006). 45 .5997. 85 8 038388 .063 8on88 BEE 05 Mo 38:85 Rama 2: 98 85:8 a B Show <75 «..MEKE 05 .2055me some E .AoweEL come So: Boo—ow 803 8:2 wEEEBE was $565me ooh: Set 88388 a 2 omen: of. 26382:»: .coeowc m< 05 e5 NOBEIoSE .3 @8355 moonEoo vote? or: 8865 30.5 new 22305 2F .33: he 882 m< 2: s 93 3 892-35 as .ecoesaa 5 N .mE Be B865 3 Ease to Ea 33.2? a? 9 2:- see 355% 92 we <75 Eéuessda 5S 33E .cBmB 55an «\wéofi 9 wane—.5 H8 aouofim m< 05 Ea NOBEEEE coming we saw—«@800 .md Semi we- 2 mm- mm. em. 2 mm. mm- 00. 0. mm- em- 2 mm- on- 9 3.- mu. 0. ms- k. 2 mm- em- 2 me. ._.>> 9.. 832.22: 46 We tested nucleotides for an effect on DNA binding by MrpC2 since it is similar to CRP- farnily transcription factors (Sun and Shi 2001b). Cyclic AMP (CAMP) allosterically controls CRP DNA binding (Kolb et a1. 1993), but the family also includes well-studied proteins like FNR with non-nucleotide effectors of DNA binding (Kiley and Beinert 1998). MrpC2 lacks certain residues that contact CAMP, so it was suggested that it might bind another nucleotide; for example, (p)ppGpp (Ueki and Inouye 2003). We measured Hislo~MrpC2 binding to fingA promoter region DNA (~101 to +25) by EMSAs and observed no effect of ppGpp (50-400 11M), cAMP (4-8 1.1M), cGMP (20-40 nM), NTPs (400 11M), or dNTPs (200 M) at concentrations designed to reflect physiological conditions (data not shown). Although we found no evidence for a nucleotide effector of DNA binding, MrpC2 might have a different type of effector. FruA is required for association of MrpC and/or MrpC2 with the fmgA promoter region in vivo The proximity of the FruA and MrpC2 binding sites in the fingA promoter region suggested that one protein might recruit the other or that the two proteins might bind cooperatively. Expression of fruA depends on MrpC2 (Figure 2.1) (Ueki and Inouye 2003), so neither transcription factor is expected to accumulate in an mrpC mutant. However, MrpC and MrpC2 accumulate normally in a fruA mutant (Nariya and Inouye 2006), yet fingA fails to be expressed (Yoder-Himes and Kroos 2006). Why are MrpC and MrpC2 insufficient to activate fingA transcription? We hypothesized that they fail to bind to the fingA promoter region in the absence of FruA. To test this hypothesis, chromatin immunoprecipitation (ChIP) with polyclonal antibodies against MrpC (which 47 also recognize MrpC2) (Nariya and Inouye 2006) was used to measure the association of MrpC and/or MrpC2 with the fingA promoter region (~101 to +155) integrated ectopically into the chromosome of wild-type or fruA mutant cells that had undergone development. DNA recovered from ChIP was subjected to PCR with primers designed to amplify the ectopic copy of the fingA promoter region. The PCR analysis revealed that the fmgA promoter region was enriched by ChIP with antibodies against MrpC relative to control antibodies for wild type, but not for the fruA mutant (Figure 2.6, top panel). Neither strain showed enrichment of rpaC coding region DNA (as a negative control) (Figure 2.6, bottom panel). We conclude that FruA is required for association of MrpC and/or MrpC2 with the fmgA promoter region in viva. Cooperative binding of MrpC2 and F ruA to the fmgA promoter region The requirement for FruA for association of MrpC and/or MrpC2 with the fingA promoter region in viva is consistent with recruitment or c00perative binding. To distinguish between these models and to test the notion that FruA directly affects binding of MrpC2 to the fingA promoter region, recombinant His-tagged FruA (FruA-His6) was mixed with Hism-MrpCZ and fingA promoter region DNA (~101 to +25) for analysis of DNA binding by EMSAs. FruA-His6 bound to the fingA promoter region weakly compared with Hislo~ MrpC2, but there was a strong enhancement of shifted complex formation when both proteins were incubated with fingA DNA (Figure 2.7A). In the presence of both proteins, two complexes were observed. The abundant lower complex (LC) co-migrated with the complexes formed by either protein alone, suggesting that the LC is a mixture of 48 complexes composed of DNA bound by Hislo-MrpCZ or FruA-His6. The upper complex (UC) was suggestive of a complex of two protein molecules bound to DNA. Wild Type fruA mutant Input Input M IgG a- MrpC MIQG a- MrpC fmgAh“ was - anal F397;}: ' rpocyia " :5 g... Figure 2.6. FruA is required for association of MrpC and/or MrpC2 with the fingA promoter region in viva. ChIP analysis of M. xanthus with the fingA promoter region (- 101 to +155) integrated ectopically in otherwise wild-type or fruA mutant backgrounds. After 18 h of development, cells were treated with formaldehyde, lysed, and crosslinked chromatin was immunoprecipitated with anti-MrpC antibodies or IgG as a control. DNA was amplified with appropriate primers for the fingA promoter region at the ectopic chromosomal site or for the rpaC coding region as a negative control. A twofold dilution series of input DNA purified from 0.25%, 0.125%, 0.0625% or 0.03125% of the total cellular extract prior to immunoprecipitation was used as a template in parallel PCRs to show that the PCR conditions were in the linear range of amplification for each primer set. Interestingly, the C-terminal DNA-binding domain of FruA was insufficient to enhance shifted complex formation in combination with MrpC2. The complexes formed by the combination of proteins were indistinguishable from those produced when FruA-DBD~ Hi53 or Hislo~MrpC2 alone was incubated with fingA promoter region DNA (Figure 2.73). We conclude that the N-terminal domain of FruA contains an important determinant for enhancement of shifted complex formation in combination with MrpC2 and the fmgA promoter region. 49 To characterize the enhanced DNA binding observed in the presence of Hislo-MrpCZ and FruA-Hi36 (Figure 2.7A), DNase I footprinting of complexes in solution was performed. Protection was observed with His lo-MrpCZ alone in the region spanning the S-bp element and the C box (Figure 2.8A), where MrpC2 binding was previously mapped by EMSAs (Figure 2.5). There were prominent hypersensitive sites in this region (Figure 2.8A), suggesting that Hislo-MrpCZ bends the DNA upon binding. The protection and hypersensitivity in this region increased when FruA-His6 was present in combination with Hislo-MrpCZ, but was not observed with FruA-His6 alone, suggesting that His 10- MrpC2 binding was increased in the presence of FI'UA-I'IlS6. DNase I footprinting of the other strand with FruA-His6 alone revealed a hypersensitive site near ~86 (Figure 2.8B), a region that was previously shown to bind FruA-DBD-Hiss (Yoder-Himes and Kroos 2006). The intensity of this hypersensitive site increased when His lo-MrpCZ was present in combination with FruA-His6, but hypersensitivity was not observed with His 10-MrpC2 alone, suggesting that FruA-His6 binding was increased in the presence of Hile-MrpCZ. As observed with the other strand (Figure 2.8A), there were protected and hypersensitive sites near the C box with Hile-MrpCZ alone and these signals were increased in the presence of both proteins (Figure 2.8B). Hypersensitive sites were observed near -77 and -70 when both proteins were present but not with either protein alone (Figure 2.8B), suggesting simultaneous binding of MrpC2 and FruA to the same DNA molecule. The 50 DNase I footprinting results demonstrate cooperative binding of FruA and MrpC2 to the fmgA promoter region, providing plausible explanations for the observed dependence of MrpC and/or MrpC2 on FruA for association with the fmgA promoter region in vivo (Figure 2.6) and for the observed enhancement of shifted complex formation in vitro (Figure 2.7A). 51 Figure 2.7. Enhancement of shifted complex formation. (A) The combination of FruA- His6 and Hislo-MrpC2 enhances complex formation. EMSAs with 32P—labeled fmgA DNA (1.2 nM) spanning from -101 to +25 with no protein, Hislo-MrpC2 (1 MM), FruA- Hi56 (3 nM) or both His 10—MrpC2 (1 nM) and FruA-His6 (3 uM) as indicated. Arrowheads indicate upper complex (UC) and lower complex (LC). (B) The combination of FruA-DBD—I-Ii33 and Hislo-MrpC2 does not enhance complex formation. EMSAs with 32P-labeled fmgA DNA (1.2 nM) spanning from -101 to +25 with no protein, Hislo- MrpC2 (1 uM), FruA-DBD-Hi38 (14 nM) or both His 10-MrpC2 (1 nM) and FruA-DBD- Hiss (14 nM) as indicated. The arrowhead and arrow indicate the complexes produced by Hislo-MrpCZ and FruA-DBD-Hi53, respectively. 52 A FruA-Hise - - + ... His1o-Mrp02 ' + - + FruA-DBD-Hi38 - - + + His1o-Mrp02 ' + ' + 53 "I qu. -i‘ w Figure 2.8. DNase I footprinting shows cooperative binding of MrpC2 and FruA to the fmgA promoter region. (A) fmgA promoter region DNA (-139 to +25) was 5'-labeled at - 139, incubated with 1 or 1.5 uM Hislo-MrpC2 (lanes l-2), or with 1.5, 3 or 4.5 uM FruA-His6 (lanes 3-5), or with 0.5, l or 1.5 uM Hislo-MrpCZ in combination with 1.5, 3 and 4.5 uM FruA-Hi36 (lanes 6-8), or with no protein (lane 9), and subjected to DNase I footprinting. (B) fmgA promoter region DNA (-101 to +25) was 5'-labeled at +25, incubated with 1.5 ptM Hislo-MrpCZ (lane 1), or with 1.5, 3 or 4.5 uM FruA-His6 (lanes 2—4), or with 0.5, l or 1.5 [1M His 10-MrpC2 in combination with 1.5, 3 or 4.5 uM FruA- His6 (lanes 5-7), or with no protein (lane 8), and subjected to DNase I footprinting. In both panels, arrows indicate sites protected from DNase I digestion and arrowheads indicate hypersensitive sites. Also, lanes G, A, T and C show sequence ladders generated by the same labeled primer used to generate the probe for DNase I footprinting. Lanes between the sequence ladders and the footprints were deleted from the images. (C) Summary of protected and hypersensitive site. 54 IS) a :6 5‘2 3:“ is “I ”if 41-1 55 FruA-His6 His1o-MrpC2 -42 C box and 5-bp ‘ ‘_ element B -41 ..4] - FruA-His,i 5-bp element and C box ‘ 1““. A . . C. lllleu—v - 8 5 GTCGGGGGTGTGCGGTGGGGAGCGAACAGTCCCACATCCCTGGCG - 4 2 SAGCCCCCAC‘IACGCCACAICCCTCGCTTGTCAGGGTfiTAGGGACCGC 56 To determine whether the binding sites for both Hislo-MrpCZ and FruA-His6 in the fmgA promoter region are important for enhanced formation of shifted complexes, EMSAs were performed with mutant DNA fragments expected to impair binding of one or the other protein. Mutations in the region from -86 to -77 greatly reduce binding of FruA- DBD-Hi33 (Yoder-Himes and Kroos 2006), and reduced the enhancement of shifted complex formation by the combination of FruA-His6 and Hislo-MrpCZ (Figure 2.9). (+5bp) WT -86 to -84 -83 to -77 -76 to -41 -63 to -60 FruA-Hise--++--++--++--++--++ Hisw-MrpCZ-+-+-+--i:.-‘+;+-+-+-+-+ Figure 2.9. Enhancement of shifted complex formation depends on binding sites for both FruA and MrpC2. EMSAs with 32P-labeled fmgA DNA ( 1.2 nM) spanning from —101 to +25, wild-type (WT) or mutant as indicated (see Fig. 2 for mutations), and Hislo-MrpCZ (1 uM) and/or FruA—His6 (3 nM) as indicated. The modified (+5 bp) —76 to -41 DNA fragment has non-fmgA sequence (CACAA) at its upstream end. UC was undetectable and LC was greatly diminished with DNA fragments bearing mutations at -86 to -84 or -83 to -77. The small amount of enhancement of LC formation 57 could be due to a small amount of FruA-Hi36 binding that is undetectable when FruA- His6 alone is incubated with DNA. To test this possibility, we attempted to eliminate the FruA—His6 binding site without impairing Hislo-MJpC2. A DNA fragment from -76 to - 41 was insufficient for His 10-MrpC2 binding (data not shown), indicating that the site required for His 10—MrpC2 binding may partially overlap the site required for FruA-His6 binding. However, adding 5 bp of non-fingA sequence (CACAA) to the upstream end allowed Hile-MrpC2 binding (Figure 2.9). No FruA-His6 binding was detected with this modified (+5 bp) -76 to —41 DNA fragment. In the presence of His 10-MrpC2 and FruA- His6, UC was undetectable and very little enhancement of LC formation was observed. These results demonstrate the importance of the FruA-I-Iis6 binding site for enhanced formation of shifted complexes. Likewise, the His 10-MrpC2 binding site is extremely important since a DNA fragment containing a mutation at -63 to -60, which eliminates detectable Hislo-MrpC2 binding, also abolished detectable enhancement of shifted complex formation (Figure 2.9). Supershift assays provided further evidence that both Hile-MrpC2 and FruA-His6 are responsible for enhanced formation of shifted complexes with fingA promoter region DNA. After incubating Hile-MrpCZ and FruA-Hi56 with fingA DNA to allow formation of complexes, purified antibodies against MrpC or FruA were added and incubation was 58 continued, followed by EMSAs. An increasing amount of anti-MrpC antibodies resulted in progressive loss of UC and LC, and appearance of supershifted complexes (Figure 2.10A). A greater amount of anti-MrpC antibodies did not result in more of the supershifted complexes. Rather, there appeared to be more unbound DNA fragment, as if the antibodies interfered with the equilibrium between bound and unbound Hislo-MrpCZ. Similar results were obtained with FruA antibodies, although inhibition of UC and LC formation was incomplete (Figure 2.10B). These results support the interpretation that enhancement of shifted complex formation involves binding of both His lo-MrpCZ and FruA-His6 to adjacent (possibly overlapping) sites upstream of the fmgA promoter, and together with our footprinting and ChIP results support a model in which FruA and MrpC2 bind cooperatively to regulate fmgA transcription during M. xanthus development. 59 Figure 2.10. Antibodies against MrijZ or FruA cause appearance of supershifted complexes and loss of UC and LC. 2—P labeled fingA promoter region DNA (1. 2 nM) spanning from -101 to +25 was incubated with Hislo-MrpCZ (1 uM) and/or FruA-Hi56 (3 nM) as indicated followed by addition of antibodies. (A) Supershift assay with an increasing amount of anti-MrpC antibodies (0.2, 0.3, 0.5 or 0.6 nM) as indicated. IgG (0.7 uM) served as a negative control. (B) Supershift assay with an increasing amount of anti-FruA antibodies (7 or 8 nM) as indicated. IgG (8 uM) served as a negative control. In both panels, arrowheads indicate the UC and LC, and a bracket indicates the supershifted complexes. 60 IgG" - - - - - - - + a-MrpC - - - -4] - FruA-Hiss - - + + + + + + + His1o-MrpC2 - + - + + + + + + oeno. IgG a-FruA FruA-Hise Hism-MrpCZ 61 DISCUSSION We have discovered that a crucial cis-regulatory element in the fingA promoter region is bound by MrpC2, but that recruitment of MrpC2 to its binding site is enhanced by FruA. Our DNA binding studies revealed cooperative binding of FruA and MrpC2 to adjacent (possibly overlapping) sites upstream of the fmgA promoter. This represents a novel mechanism of gene regulation since recruitment of another transcription factor (MrpC2) by a response regulator (FruA) has not been observed previously, despite the prevalence of two-component signal transduction systems, especially in bacteria. Our preliminary results, described below, indicate that several other promoter regions that depend on C- signaling for activation are cooperatively bound by FruA and MrpC2. Since MrpC2 is a proteolytic fragment of MrpC, and these proteins function not only as transcription factors, but also in the regulation of PCD, it appears that cooperative binding of FruA and MrpC2 facilitates the coordination of multiple signaling pathways to ensure proper control of gene expression and cell fate during M. xanthus development. Cooperative binding of MrpC2 and FruA appears to be a conserved mechanism of gene regulation in response to C-si gnalin g during M. xanthus development. The cis-regulatory element to which MrpC2 binds in the fingA promoter region includes a 5-bp element and a C box. These two sequences are similarly arranged immediately upstream of other C- signal-dependent promoters and are important for promoter activity (Viswanathan and Kroos 2003, Yoder and Kroos 2004b, Loconto et al. 2005, Viswanathan et al. 2006a), suggesting that MrpC2 may bind to these sites. Indeed, in the promoter region of the operon identified by Tn5 lac £24499, MrpC2 binds near a 5-bp element and it appears to 62 bind cooperatively with FruA (S. M. and L. K., unpublished data). In the promoter region of the dev operon, whose products are important for sporulation (Thony-Meyer and Kaiser 1993, Boysen et al. 2002, Viswanathan et al. 2007b), MrpC2 binds to a region that includes a 5-bp element followed 3 or 7 bp downstream by two C-box-like sequences, and it appears to bind cooperatively with FruA (S. M., P. Viswanathan, and L. K., unpublished data). In the promoter region of the gene identified by Tn5 lac £24403, MrpC2 binds to a region that includes two 5-bp elements in inverted orientation, and it appears to bind cooperatively with FruA (J. Lee, S. M., and L. K., unpublished data). Our preliminary studies, taken together with the evidence presented here for fmgA, indicate that cooperative binding of MrpC2 and FruA is a conserved mechanism of C-signal- dependent gene regulation. Cooperative binding of MrpC2 and FruA to promoter regions of C-signal-dependent genes represents a novel mechanism of gene regulation. Typically, DNA-binding response regulators are phosphorylated by an HPK and this modification enhances DNA binding (West and Stock 2001). The bound response regulator recruits RNA polymerase to the promoter or facilitates another step during transcription initiation. Phosphorylation of FruA’s N-terminal regulatory domain may relieve an inhibitory effect on its C- terrninal DNA-binding domain, since FruA-DBD-Hi33 appears to bind with higher affinity than (presumably unphosphorylated) full-length FruA-Hi36 to the fmgA promoter region (Fig. 6) and other promoter regions (Viswanathan et al. 2007b) (S. M. and L. K., unpublished data). We found that FruA is required for association of MrpC and/or MrpC2 with the fingA promoter region in vivo (Figure 2.6). This presumably explains why fingA 63 expression is abolished in a fruA mutant (Yoder-Himes and Kroos 2006). FruA and/or MrpC2 probably interact with RNA polymerase at the fingA promoter. The two proteins occupy a location typical for Class I activators, which function by contacting the C- terminal domain of the on subunits of RNA polymerase (Barnard et al. 2004). The detailed mechanism of cooperative binding of MrpC2 and FruA to the fmgA promoter region remains to be explored. The binding sites of the two proteins may partially overlap since a 7-bp mutation at -83 to ~77 impairs FruA-DBD-HiS3 binding (Yoder-Himes and Kroos 2006) and since DNA upstream of -76 is required for His")- MrpC2 binding (data not shown). The two proteins may interact with opposite faces of the DNA in a region of overlap, analogous to certain homeodomain proteins, which bind DNA cooperatively (Tullius 1995, Passner et al. 1999). As for some homeoprotein-DNA complexes, cooperativity might depend not only on protein-protein interactions but on bending of the DNA by one or both proteins. Binding of either MrpC2 or FruA alone to the fmgA promoter region produced DNase I hypersensitivity indicative of DNA bending, and the combination of proteins increased the intensity and number of hypersensitive sites (Figure 2.8). While these results demonstrated cooperative binding, we did not observe much protection from DNase I digestion, suggestive of limited occupancy. Likewise, EMSAs clearly showed that the combination of proteins enhances formation of shifted complexes (Figure 2.7A) and that this depends on sequences important for binding of each protein (Figure 2.9), consistent with cooperative binding, yet the predominant shifted complex, LC, co-migrated with complexes produced by either protein alone. This suggests limited co-occupancy by MrpC2 and FruA, although we 64 cannot rule out the possibility that LC is a mixture of binary (one protein bound to DNA) and ternary (both proteins bound to DNA) complexes since the ternary complex might migrate unexpectedly due to DNA bending. The enhancement of shifted complex formation by MrpC2 and FruA requires the N-terminal regulatory domain of FruA (Figure 2.7B), which is believed to be phosphorylated by one or more HPKs in M. xanthus (Ellehauge et al. 1998). Treatment of FruA-His6 (purified from E. coli and therefore conceivably phosphorylated to a small extent) with phosphatase from bacteriophage A did not diminish its ability to enhance formation of shifted complexes (data not shown). It seems likely that the N-terminal domain of FruA interacts directly with MrpC2, but further studies will be needed to distinguish this model from the possibility that FruA’s N-terminal domain alters the DNA structure in a way that facilitates MrpC2 binding. No enhancement of shifted complex formation was observed when the FruA and MrpC2 binding sites were on separate DNA fragments (only one of which was 32P-labeled in each of two separate experiments) (data not shown). Although the detailed mechanism of cooperative binding is unknown, it is worth exploring because response regulators like FruA and CRP-family transcription factors like MrpC are abundant in bacterial signaling and gene regulatory networks, and cooperative binding between such proteins could provide a general mechanism to achieve tight combinatorial control of target genes. The regulation of fmgA by the combination of MrpC2 and FruA constitutes a coherent feed-forward loop. Such loops, in which one transcription factor (MrpC2) positively regulates another (FruA), and the two factors both positively regulate target genes, are 65 common in regulatory networks (Milo et al. 2002, Shen-Orr et al. 2002, Mangan and Alon 2003). They allow filtering out of noise in input stimuli, rapid response to step—like stimuli in one direction (e.g., off to on), delayed response to steps in the opposite direction (e. g., on to off), and delay of target gene expression until both transcription factors reach a sufficient concentration (Mangan et al. 2003). These characteristics could have obvious benefits for regulation of fmgA and other genes in response to starvation, C- signaling, and other signals via MrpC2 and FruA. Combinatorial regulation of gene expression is common in bacteria and eukaryotes because it allows multiple signals to control individual genes (Barnard et al. 2004, Stathopoulos and Levine 2005). Cooperative interactions between activators is one of several mechanisms for achieving combinatorial control in bacteria; however, not many examples (and none involving a response regulator) have been reported previously, perhaps due to evolutionary constraints it places on the activators (Barnard et al. 2004). For transcription factors like MrpC and FruA that are devoted to a developmental program, such evolutionary constraints may be tolerable. On the other hand, both MrpC and FruA probably interact with multiple protein partners, in addition to interacting with DNA, and possibly with each other and RNA polymerase. It has been proposed that distinct HPKs phosphorylate FruA in response to an early (unidentified) signal and later in response to C-signal (Ueki and Inouye 2003) (Figure 2.1). This model can explain how different FruA-dependent genes exhibit different levels of dependence on C-signaling, if in the absence of C-signaling the level of 66 phosphorylated FruA supports full (e. g., dofA), partial (e. g., fmgA), or no expression due to differential affinity for binding sites in promoter regions. It will be interesting to test whether C-signal-independent genes like dofA are not only directly regulated by FruA (Ueki and Inouye 2005a) but also by cooperative binding of MrpC2. MrpC is phosphorylated by an STPK cascade, presumably in response to an unknown signal during growth, inhibiting accumulation of MrpC and MrpC2 (Nariya and Inouye 2005). Starvation conditions may remove the signal (Figure 2.1), allowing MrpC and MrpC2 to accumulate. Recently, it was shown that the EspA signal transduction pathway influences the MrpC and MrpC2 concentrations (Higgs et al. 2008), presumably providing another link to starvation (Figure 2.1), although the exact signal to which EspA responds is unknown. Also, MrpC was shown to interact with the toxin MazF, inhibiting PCD (N ariya and Inouye 2008) (Figure 2.1). On the other hand, MrpC appears to directly activate mazF transcription. Whether MrpC2 differs from MrpC in either of these activities is important to test. The concentrations of MrpC and its phosphorylated or cleaved forms, and their interactions with MazF, and at different promoters, may determine the fate of cells in a developing population of M. xanthus. Nariya and Inouye (Nariya and Inouye 2008) suggested that the position of an individual cell in the cell cycle at the time of nutrient depletion might determine its fate. Alternatively, by analogy with many cell fate decisions in bacteria (Dubnau and Losick 2006, Smits et al. 2006) and eukaryotes (Ferrell 2002), we suggest that the MrpC positive autoregulatory loop could be a source of bistability in the developing cell population. 67 This model predicts heterogeneity in the MrpC concentration in cells undergoing development. The concentrations of MrpC and MrpC2 also influence the pace of aggregation and the morphology of fruiting bodies. Mutants defective in the STPK cascade (that leads to phosphorylation of MrpC) or EspA exhibit accelerated aggregation (Nariya and Inouye 2005, I-Iiggs et al. 2008). These mutants make fairly normal spore numbers. Whether MazF-mediated PCD is aberrant in these mutants and whether this contributes to the disorganized appearance of fruiting bodies and loss of coordination between aggregation and sporulation (i.e., spore formation outside of fruiting bodies) are intriguing questions. Commitment to form a spore has been hypothesized to involve induction of genes at the $27536 locus, which in turn depends on induction of the dev operon (Kroos 2007). Since dev appears to be regulated by cooperative binding of MrpC2 and FruA (S. M., P. Viswanathan, and L. K., unpublished data), we propose that commitment to sporulation is governed by these key transcription factors. MrpC is a major hub in the regulatory network, linked extensively to starvation (Figure 2.1). Its direct involvement in commitment to sporulation might couple persistent starvation to the decision to form a spore. FruA is likewise a major hub in the regulatory network. Phosphorylation of FruA in response to short-range C-signaling might contribute positional information (i.e., cell alignment in the nascent fruiting body) to the decision to sporulate, ensuring that spores form within fruiting bodies (Kaiser 2003, Sogaard-Andersen et al. 2003). Other potential inputs via FruA include phosphorylation in response to another signal(s) (Jelsbak et al. 2005) and regulation at the transcriptional level. In addition to direct regulation by 68 MrpC2 (Ueki and Inouye 2003), at least four other inputs directly or indirectly control fruA transcription; SasR (responsive to A-signal) (Guo et al. 2000), chA (Nielsen et al. 2004), CarD (Penalver-Mellado et al. 2006), and DevT (feeding back positively from the dev operon) (Boysen et al. 2002). Commitment to sporulation may also be governed by a third activator of dev transcription, LadA (Viswanathan et al. 2007b). This LysR-type transcription factor likely responds to a signal and, unusually, it acts positively from a site downstream of the dev promoter, perhaps by counteracting negative regulatory elements. Combinatorial regulation of dev by at least three signal-responsive transcription factors and a regulatory region spanning more than 1 kb resembles regulation of developmental genes in multicellular eukaryotes (Stathopoulos and Levine 2005, Viswanathan et al. 2007a, Viswanathan et al. 2007b). Integration of environmental, cell—to—cell, and intracellular signals by transcription factors to control gene expression and cell fate is crucial for multicellular organisms, especially during development. We have discovered that during M. xanthus development the signal integration potential of MrpC is combined with that of FruA. Cooperative binding, and a coherent feed-forward loop design, create a powerful regulatory circuit to achieve correct temporal and spatial expression of target genes. Examples of spatiotemporal differentiation of bacteria in architecturally complex biofilms continue to emerge (Vlamakis et al. 2008), so it would be surprising if the design principles discovered in M. xanthus are not utilized in other bacterial communities, as well as in multicellular eukaryotes. 69 Chapter 3: Combinatorial regulation by a novel arrangement of FruA and MrpC2 transcription factors during Myxococcus xanthus development The work in this chapter will be submitted to the Journal of Bacteriology in September 2008. 70 ABSTRACT Myxococcus xanthus is a Gram-negative soil bacterium that undergoes multicellular development upon nutrient limitation. Intercellular signals control cell movements and regulate gene expression during the developmental process. C-signal is a short-range signal essential for aggregation and sporulation. C-signaling regulates the fmgA gene by a novel mechanism involving cooperative binding of the response regulator FruA and the transcription factor/antitoxin MrpC2. Here, we demonstrate that regulation of the C- signal—dependent fingBC operon is under similar combinatorial control by FruA and MrpC2, but the arrangement of binding sites is different than in the fmgA promoter region. MrpC2 was shown to bind to a crucial cis-regulatory sequence in the fmgBC promoter region. FruA was required for MrpC and/or MrpC2 to associate with the fmgBC promoter region in vivo, and expression of an fmgB-lacZ fusion was abolished in a fruA mutant. Recombinant FruA was shown to bind to an essential regulatory sequence located slightly downstream of the MrpC2-binding site in the fmgBC promoter region. Full-length FruA, but not its C-terminal DNA-binding domain, enhanced the formation of complexes with fingBC promoter region DNA, when combined with MrpC2. This effect was abolished with fmgBC DNA fragments having a mutation in either the MrpC2- or FruA-binding site, indicating that both proteins must bind to DNA in order to enhance complex formation. These results are similar to those observed for fingA, where FruA and MrpC2 bind cooperatively upstream of the promoter, except in the fmgA promoter region the FruA-binding site is located slightly upstream of the MrpC2-binding site. Cooperative binding of FruA and MrpC2 appears to be a conserved mechanism of gene 71 regulation that allows a flexible arrangement of binding sites, and coordinates multiple signaling pathways. 72 INTRODUCTION Myxococcus xanthus is a rod-shaped bacterium that glides on solid surfaces, forming a single-species biofilm that provides an attractive model to study how signaling couples gene expression to environmental and cellular cues (Whitworth 2008). M. xanthus cells in the biofilm grow and divide when nutrients are available, but upon starvation, a multicellular developmental process ensues, during which cells move into aggregates and form mound-shaped structures called fruiting bodies. Approximately 105 cells participate in forming a fruiting body, in which a portion of the cells differentiate into dormant, stress-resistant, spherical spores. Other cells undergo programmed cell death (Nariya and Inouye 2008) or autolysis caused by siblings in the developing biofilm (Wireman and Dworkin 1977, O'Connor and Zusman 1988), and some cells remain outside of fruiting bodies as peripheral rods (O'Connor and Zusman 1991). These fates are met by different proportions of cells in the biofilm, depending on genetic and environmental factors (O'Connor and Zusman 1988, Berleman and Kirby 2007). The spores in a fruiting body can germinate and resume growth and division when nutrients become available. Signals act at different times during the developmental process to coordinate cell behavior and determine cell fate. Nutrient limitation causes a stringent response that results in production of (p)ppGpp and the induction of early developmental genes (Harris et al. 1998). A mixture of amino acids and peptides, known as A-signal, is generated by secreted proteases and is believed to allow quorum sensing (Kuspa et al. 1992b). A- signal-dependent genes are expressed and cells alter their pattern of movement so that aggregates begin to form. Subsequent gene expression, and the maturation of aggregates 73 into spore-filled fruiting bodies, depends on C-signaling, which is mediated by the product of the csgA gene (Shimkets et al. 1983). ngA is associated with the outer membrane of the cell, where it is processed by a secreted protease to a 17 kDa form that appears to act as a short-range signal (Kim and Kaiser 1990d, Lobedanz and Sogaard- Andersen 2003). C-signal transduction requires cell-alignment (Kim and Kaiser 1990b) and possibly end-to-end contact between cells (Sager and Kaiser 1994), so it communicates positional information. Cells become aligned as aggregates transform into nascent fruiting bodies and the resulting high level of C-signaling has been proposed to trigger expression of genes required for sporulation (Sager and Kaiser 1993). Indeed, the expression of C-signal-dependent genes that are important for sporulation is restricted to nascent fruiting bodies (Sager and Kaiser 1993, Julien et al. 2000), and many studies support a model in which an increasing level of C-signaling controls gene expression to coordinate aggregation and sporulation during development (Kim and Kaiser 1991, Li et al. 1992, Gronewold and Kaiser 2001, Kruse et al. 2001). How does C-signaling regulate expression of target genes? FruA plays a key role in the C-signal transduction pathway (Ogawa et al. 1996, Ellehauge et al. 1998). It is similar to response regulators of two-component signal transduction systems and is believed to be phosphorylated in its N-terminal regulatory domain in response to C-signal and perhaps other signals (Ellehauge et al. 1998, Jelsbak et al. 2005), but the cognate histidine protein kinase(s) has not been identified. Presumably, phosphorylation enhances DNA binding by the C-terminal domain of FruA, which is similar to that of the NarUFixJ subfamily of response regulators (West and Stock 2001). The C-terminal domain of FruA has been 74 shown to bind to sites in the promoter regions of developmental]y-regulated genes that fail to be expressed in fruA mutant cells, suggesting that FruA is a transcriptional activator (Ueki and Inouye 2005a, Ueki and Inouye 2005b, Yoder-Himes and Kroos 2006). Recently, FruA was shown to bind cooperatively with MrpC2 to the promoter region of the C-signal-dependent fmgA (EruA- and MrpC2-regulated gene A) gene (Mittal and Kroos 2008), revealing a novel mechanism of combinatorial control, as cooperative binding of a response regulator (FruA) and a distinct transcription factor (MrpC2) had not been observed previously. MrpC2 is a smaller form of MrpC (Ueki and Inouye 2003), which is similar to the cyclic AMP receptor protein (CRP) family of transcriptional regulators (Sun and Shi 2001b). MrpC is expressed during vegetative growth and is phosphorylated by a cytoplasmic serine/threonine protein kinase (STPK) called Pknl4 (Nariya and Inouye 2005, Nariya and Inouye 2006). Pan4 is in turn phosphorylated by a membrane STPK called Pkn8. Phosphorylation of MrpC by the Pkn8/Pknl4 cascade results in weaker binding of MrpC to DNA and also appears to inhibit proteolytic cleavage of MrpC to MrpC2 (Nariya and Inouye 2006), which lacks the 25 N-terminal residues of MrpC (Ueki and Inouye 2003). The STPK cascade is counteracted by an unknown mechanism early in development, allowing MrpC and MrpC2 concentrations to rise. MrpC2 binds to DNA with higher affinity than MrpC (Nariya and Inouye 2006), and appears to play a key role as a transcriptional activator during development. Recently, MrpC was shown to function as an antitoxin by interacting directly with the toxin MazF, an mRNA interferase that mediates programmed cell death during development (Nariya and Inouye 2008). MrpC 75 alsr the Elf DO (ii M also binds to the mazF promoter region and activates expression. Binding of MrpC2 to the mazF promoter region and MazF has not been tested. The dual functions of MrpC, and possibly MrpC2, as an antitoxin and a transcription factor make it an important determinant of cell fate. The finding that MrpC2 and FruA bind cooperatively to crucial cis-regulatory sequences upstream of the fmgA promoter suggests that starvation and other signals that regulate MrpC2 are integrated with positional information via short- range C-signaling that leads to phosphorylation of FruA (Mittal and Kroos 2008). This novel mechanism of combinatorial control was predicted to be conserved because similar cis-regulatory sequences have been found upstream of other developmentally-regulated M. xanthus promoters (Fisseha et al. 1999, Viswanathan and Kroos 2003, Yoder and Kroos 2004b, Srinivasan and Kroos 2004, Loconto et al. 2005, Viswanathan et al. 20063, Viswanathan et al. 2007a). The promoter region of a putative operon (named herein fingBC for EruA- and MrpC2- regulated genes B and Q) at the (24499 locus in the M. xanthus chromosome has cis- regulatory sequences similar to those bound by MrpC2 in the fingA promoter region. The fmgBC operon was identified by an insertion of the transposon Tn5 lac into fingC (Kroos et al. 1986). FmgB and FmgC are similar to reductase and oxidase components, respectively, of bacterial cytochrome P-450 systems, which typically are involved in catabolism or anabolism of unusual compounds (Fisseha et al. 1999). M. xanthus DNA upstream of fmgBC was cloned, a putative transcriptional start site was mapped, and the region from -100 to +50 was shown to encompass the promoter (Fisseha et al. 1999, Yoder and Kroos 2004b). Expression from the fmgBC promoter was reduced in a csgA 76 mut. “his sigh Cllil [“0 the pro b0.‘ the fm IE) mutant but was restored upon co-development of the csgA mutant with wild-type cells, which supply C-signal, demonstrating that promoter activity is partially dependent on C- signaling (Kroos and Kaiser 1987, Fisseha et al. 1999). Mutational analysis identified critical cis-regulatory sequences at -71 to -45 upstream of the promoter (Yoder and Kroos 2004b). This region contains two C boxes (consensus CAYYCCY; Y means C or T) and two 5-bp elements (consensus GAACA) (Figure 3.1), which are sequence motifs found in the promoter regions of several developmentally-regulated genes (Fisseha et al. 1999, Viswanathan and Kroos 2003, Yoder and Kroos 2004b, Srinivasan and Kroos 2004, Loconto et al. 2005, Viswanathan et al. 2006a, Viswanathan et al. 2007a). In the fingA promoter region, between -63 and -46, a 5-bp element is located 6 bp upstream of a C box, and this region is bound by MrpC2, while FruA binds cooperatively to a site located slightly upstream (Mittal and Kroos 2008). Here, we report that MrpC2 and FruA bind to sequences between -71 and -45 upstream of the fingBC promoter, but the arrangement of binding sites is the reverse of that found in the fingA promoter region. Nevertheless, the association of MrpC and/or MrpC2 with the fmgBC promoter region in vivo required FruA. Furthermore, there appeared to be cooperative binding of MrpC2 and FruA to fmgBC promoter region DNA in vitro. Our results demonstrate combinatorial control by MrpC2 and FruA at a second promoter, and reveal surprising flexibility in the arrangement of the binding sites. 77 MATERIALS AND METHODS Bacterial strains and plasmids. Strains and plasmids used in this study are listed in Table 3.1. Growth and development. Escherichia coli BL21(DE3) containing plasmids were grown at 370C in Luria-Bertani (LB) medium (Sambrook et al. 1989) containing 200 pg ampicillin per ml. M. xanthus strains were grown at 320C in C11“ (1% Casitone, 10 mM Tris-HCI [pH 8.0], 1 mM KH2P04-K2HPO4, 8 mM MgSO4, [final pH 7.6]) medium (Hodgkin and Kaiser 1977) or on C'IT agar (1.5%) plates. When required, 40 ug kanamycin sulfate per ml was added. Fruiting body development was performed on TPM agar (1.5%) plates (10 mM Tris-HCl [pH 8.0], 1 mM KH2PO4-K2HPO4, 8 mM MgSO4, [final pH 7.6] as described previously (Kroos et al. 1986). Construction of M. xanthus strains and determination of lacZ expression during development. Strains containing pREGl727 or its derivatives integrated at the Mx8 phage attachment site, attB, were constructed by electroporation (Kashefi and Hartzell 1995) of M. xanthus, and transformants were selected on CTT agar plates containing kanamycin sulfate. Transformants were screened on TPM agar plates containing 40 ug of 5-bromo-4-chloro-3-indolyl-B-D-galactopyranoside per ml, in order to avoid rare transformants with unusual developmental lacZ expression (Viswanathan and Kroos 2003). Three transformants were chosen for further analysis and B-galactosidase activity was measured as described previously (Kroos et al. 1986). 78 Preparation of DNA fragments. DNA fragments spanning the fingBC promoter region from -104 to -29 were generated by PCR using wild-type or mutant plasmid (Table 3.1) as the template and the oligonucleotide primers 5’-CGCGAGGAGATI‘GCG'ITCATAC- 3' (for -104) and 5'- GAGGAATGGGCCGGAAGTTC-3' (for -29). For EMSAs, 32P- labeled DNA was synthesized by PCR after labeling the primers with [7'32P]ATP using T4 polynucleotide kinase (New England Biolabs) and the DNA fragment was purified after 15% PAGE (Sambrook et al. 1989). EMSAs. EMSAs were performed as described previously (Yoder-Himes and Kroos 2006), except that binding reaction mixtures were incubated at 25°C for 15 min. DNA-affinity chromatography. An fmgBC DNA fragment (-104 to -29) was synthesized by PCR with a 5’-biotin label at -104, bound to streptavidin beads, and DNA- affinity chromatography was performed with the AS fraction as described previously (Viswanathan et al 2007b). Preparation of Hislo-MrpCZ, FruA-His6 and FruA-DBD-Hiss. Recombinant proteins were expressed in E. coli and purified as described previously (Yoder-Himes and Kroos 2006, Nariya and Inouye 2006, Mittal and Kroos 2008). ChIP. M. xanthus strains MDY1727.DZF1, MSM4499.DZFl and MSM4499.FA were used for ChIP as described previously (Yoder-Himes and Kroos 2006). The primers used 79 for PCR of the fmgBC promoter region integrated ectopically were 5'- CT GCCAGGAATTGGGGATC-3' (upstream primer in the vector) and 5’— CGGATCCAGCGGGTGAGGTCGACGACG-3' (downstream primer with its 5' end at +50 of fmgBC). The primers used for PCR of the vector alone integrated ectopically were the same upstream primer as above and 5'- CGGGCCATCCGCCAGTGG-3' (downstream primer in the vector). The primers used for PCR of the rpoC coding region were described previously (Yoder-Himes and Kroos 2006). 80 TABLE 3.1 Strains and plasmids used in this study. Strain or Description Reference or plasmid Source E. coli BL21(DE3) F' ompT hstB(rB‘ mB‘) gal dcm with DE3, a Novagen it prophage carrying the T7 RNA polymerase gene SMhisMrpC2 BL21(DE3) containing pET16b/His10-MrpC2 (2133?? and Kroos SMFruAhis BL21(DE3) containing pETl 1km/FruA-His6 (21214631 and Kroos EDYFruA BL21(DE3) containing pETl lalFDBD-Hg (Yoder-Himes and Kroos 2006) M. xanthus DK1622 Wild type (Kaiser 1979) DK4499 Tn5 lac Q4499 (Kroos et al. 1986) MDY1727.DZF1 sglAI atthszEG1727 (Yoder-Himes and Kroos 2006) MSM1727.FA sglAI fruA::TnV 0786 atthszEG1727 This work MSM4499.DZFl sglAI atthzpDYSI This work MSM4499.FA sglAI frquzTnV 0786 attB::pDY51 This work Plasmids pETl la/FDBD- pETl 1a with a gene encoding FruA-DBD-I-Iiss (Ueki and Inouye H3 under control of a T7 RNA polymerase 2005a) promoter PET16b/Hi310' pET16b with a gene encoding His 10-MrpC2 (Nariya and MrpC2 under control of a T7 RNA polymerase Inouye 2006) promoter PET] 1km/FruA- pETl 1km with a gene encoding FruA-His6 3' Inouye H‘S6 under control of a T7 RNA polymerase promoter pREGl727 Apr Kmr Pl-inc attP ’lacZ (Fisseha et al. 1996) pDYSl pGEM7Zf with fingBC DNA from -100 to +50 (Yoder and Kroos generated by PCR using pDY100 as template 2004b) pDY100 pGEM7Zf with fmgBC DNA from -218 to +50 (Yoder and Kroos 2004b) pDYl33 pDYIOO with GCCGC-to-TAATA mutation at (Yoder and Kroos —81 to —77 2004b) pDY129 pDY100 with GGAC-to-TTCA mutation at - (Yoder and Kroos 71 to —68 2004b) pDY127 pDYlOO with ACCA-to-CAAC mutation at - (Yoder and Kroos 67 to —64 2004b) 81 Table 3.1 (cont’d) pDY125 pDY100 with CCGG-to-AA'IT mutation at (Yoder and Kroos —63 to —60 2004b) pDY49 pDY100 with TCATTC-to-GACGGA (Yoder and Kroos mutation at —59 to —54 2004b) pDY121 pDY100 with CCTTC-to-AAGGA mutation at (Yoder and Kroos —53 to -49 2004b) pDY47 pDY100 with GAAC-to-TCCA mutation at — (Yoder and Kroos 48 to -45 2004b) pDY117 pDY100 with C-to-A mutation at -37 (Yoder and Kroos 2004b) pDY45 pDY100 with CATTCCT-to-ACGGAAG (Yoder and Kroos mutation at —36 to 30 2004b) 82 RESULTS An insertion in fmgC reduces spore formation. M. xanthus strain DK4499 contains Tn5 lac £24499 inserted in fmgC, which was predicted previously to encode an oxidase of a cytochrome P—450 system (Fisseha et al. 1999). fingC corresponds to MXAN4127 in the annotation of the genomic sequence (Goldman et al. 2006). Only 59 bp upstream of fmgC is fingB (MXAN4126), which was predicted previously to code for a reductase likely to function in the same P-450 system as FmgC, although the substrate and products of the system are unknown (Fisseha et al. 1999). The short distance between fmgB and fmgC, and the finding that their products are likely components of a P-450 system, suggested that the two genes might be co-transcribed. In agreement, 5’-deletion analysis and mapping of an mRNA 5’ end, located a promoter upstream of fmgB capable of driving expression of lacZ during development similar to that observed for DK4499 containing Tn5 lac £24499 (Fisseha et al. 1999). The gene upstream of fmgB is in the opposite orientation (Goldman et al. 2006). The gene downstream of fmgC is in the same orientation, but is separated from the end of fmgC by an intergenic region of at least 243 bp and is predicted to encode a transposase, so it is unlikely to be co-transcribed with the putative fingBC operon. M. xanthus DK4499 bearing Tn5 lac £24499 aggregated normally under conditions that induce development, but the number of heat- and sonication-resistant spores that were able to germinate and form a colony was 6-fold lower than observed for wild-type DK1622. The reduced sporulation of DK4499 is likely due to loss of FmgC, although we cannot rule out an effect of the Tn5 lac insertion on expression of fmgB (e.g., due to 83 altered mRNA stability) or a gene downstream of fmgC (i.e., if transcription from the fmgBC promoter normally reads through a downstream gene). Nevertheless, our results suggest that transcription from the fingBC promoter is important for sporulation. MrpC2 binds to a key cis-regulatory sequence in the fmgBC promoter region. Mutational analysis of the fmgBC promoter region was performed previously (Yoder and Kroos 2004b) and showed that sequences upstream of the promoter are important for its activity (Figure 3.1). These regulatory sequences include two 5-bp elements and two C boxes, which are found in the promoter regions of several developmentally-regulated genes (Fisseha et al. 1999, Viswanathan and Kroos 2003, Yoder and Kroos 2004b, Srinivasan and Kroos 2004, Loconto et al. 2005, Viswanathan et al. 2006a, Viswanathan et al. 2007a). To identify putative transcription factors, we performed electrophoretic mobility shift assays (EMSAs) with a DNA fragment from the fmgBC promoter region and partially-purified DNA-binding proteins (AS fraction) from M. xanthus cells that had undergone 12 h of development, since fmgBC is expressed at this time (Kroos et al. 1986). A single shifted complex was observed with a DNA fragment spanning from -104 to -29, but no complex was observed when the DNA fragment contained a mutation in the sequence from -67 to -64 (Figure 3.1). Since this mutation was shown previously to eliminate fmgBC promoter activity in vivo (Yoder and Kroos 2004b), these results showed that a protein in the AS fraction binds to a crucial cis-regulatory sequence upstream of the fingBC promoter. 84 To purify the putative activator protein from the AS fraction, DNA-affinity chromatography was performed with the fmgBC DNA fragment (-104 to -29). The major species after purification was approximately 30 kDa in size (Figure 3.2A). The affinity- purified protein (APP) generated a shifted complex of similar mobility as observed with the AS fraction when the fmgBC DNA fragment with the wild-type sequence was used in EMSAs, and no complex was observed with the APP and the mutant (-67 to -64) fmgBC promoter region (Figure 3.2B). It appeared that APP contained the putative activator protein from the AS fraction. To identify the putative activator protein, the APP was subjected to mass spectrometry analysis after protease digestion. The peptide sequences primarily matched MrpC, a protein that is about 30 kDa in size, consistent with the size of the major species in the APP (Figure 3.2A). MrpC is similar to CRP-family transcription factors and is an essential for M. xanthus development (Sun and Shi 2001b). MrpC2, a shortened form of MrpC that lacks the 25 N-terminal residues, is produced during development and was identified in an AS fraction previously by DNA-affinity chromatography with the fruA promoter region (Ueki and Inouye 2003). We infer that MrpC2 in the AS fraction and in the APP is responsible for the shifted complex we observed with fmgBC promoter region DNA. 85 1 70 GATAG 5-bp element C box 5-bp element C box -81 GCCGQ'I'E-CfiGGAmCGGfi'CA'lTCbCTTCmTTCCGG‘JATTCCT -3o .1 1H t 1.1.33 H Figure 3.1. Effects of mutations on fingBC promoter activity in vivo and on DNA binding in vitro. The top part shows a summary of mutational effects on developmental fingB-lacZ expression (Yoder and Kroos 2004). The wild-type fmgBC upstream sequence is alternately boxed or underlined to indicate changed sequences, which are shown below the downward arrows. The number beneath each mutant sequence indicates the maximum B-galactosidase activity during development, expressed as a percentage of the maximum activity observed for the wild-type promoter. The bottom part shows EMSAs performed with 32P-labeled fmgBC DNA (12 nM) spanning from - 104 to -29 and proteins in the AS fraction (0.7 jug/pl). The arrow indicates the shifted complex produced by incubating the wild-type (WT) DNA fragment with the AS fraction. No complex was observed with a DNA fragment bearing the indicated mutation at -67 to -64. 86 7—105 3—78 :_55 WTmutant » APP - - + - - + —45 AS-+--+- 1—34 —17 Figure 3.2. DNA-affinity purification of protein that binds to the fmgBC promoter region. (A) SDS—PAGE of protein purified from the AS fraction using fingBC DNA (-104 to - 29). The arrow indicates the major species in the affinity—purified protein (APP) after staining with silver. Numbers indicate the migration positions of molecular weight (kDa) standards. (B) EMSAs with 32P-labeledfmgBC DNA (12 nM) spanning from -104 to -29 and proteins in the AS fraction or the APP. Arrowheads indicate the shifted complexes produced with the wild-type (WT) DNA fragment. No complex was observed with a DNA fragment bearing the ACCA to CAAC mutation at —67 to -64 (mutant). To confirm that MrpC2 binds to the fmgBC promoter region fragment, N-terminally His- tagged MrpC2 (Hile-MrpC2) was expressed in E. coli and purified. Hislo-MrpCZ displayed a similar pattern of binding to wild-type and mutant fmgBC DNA fragments as the AS fraction (Figure 3.3). The slower migration of the complex produced by His 10- MrpC2, as compared with the complex produced by the AS fraction, is presumably due to the 10 His residues plus 8 additional residues present in the Hislo—MrpC2 fusion protein. The mutation from —67 to -64 that resulted in loss of shifted complex formation with the 87 AS fraction (Figure 3.1) and the APP (Figure 3.2B), also caused loss of shifted complex formation with His 10—MrpC2 (Figure 3.3). This mutation includes one base pair of a 5-bp element (Figure 3.1); however, an adjacent mutation at -71 to -68, which changes the remaining four base pairs of the 5-bp element, did not impair formation of shifted complexes with the AS fraction or with Hisjo-MrpC2 (Figure 3.3). Likewise, none of the other mutations between -63 and -30 impaired complex formation. The mutation from - 81 to -77 resulted in diminished formation of the complex that we believe contains MrpC2, by the AS fraction, and the appearance of a novel shifted complex. The novel complex appears to be due to an unknown protein in the AS fraction that is capable of binding to this mutant fmgBC DNA fragment, since purified Hislo-MrpCZ did not show this effect. 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