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This is to certify that the

dissertation entitled

Characterization of regulatory mechanisms of cell-cell
interaction-dependent genes of Myxococcus xanthus

presented by

Tong Hao

has been accepted towards fulfillment
of the requirements for

PhD. Biochemistry

degree in

 

 

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Date ‘j/iozol

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6/01 cJCIRC/DateOuo.p65-p.15

 

CHARACTERIZATION OF REGULATORY MECHANISMS OF CELL-CELL
INTERACTION-DEPENDENT GENES OF MYX 0C CC C US XAN T H US

By

Tong Hao

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Department of Biochemistry and Molecular Biology

2001

ABSTRACT

CHARACTERIZATION OF REGULATORY MECHANISMS OF CELL-CELL
INTERACTION-DEPENDENT GENES OF MYXOCOC C US XAN 771 US

By

Tong Hao

Myxococcus xanthus is a gram-negative, rod-shaped soil bacterium that undergoes
multicellular development in response to nutrient limitation. Upon starvation, many
thousands of cells move in a coordinated fashion into aggregation centers and construct
multicellular structures known as fiuiting bodies, in which individual cells undergo
morphological and physiological changes to form dormant ovoid-shaped spores. The
progression through fruiting body formation and sporulation is highly coordinated and
involves at least five extracellular signaling interactions. These five signaling interactions
are also needed for the proper expression of developmental genes.

One of the extracellular signals is C-signaling, which is required for aggregation
and sporulation. C-signaling mutants initiate development normally but fail to form
compact fiuiting bodies and spores. Of the genes that have been examined, almost every
one expressed at six hours or later in development exhibits reduced or abolished
expression in C-signaling mutant cells.

To understand how extracellular C-signaling regulates gene expression during
development, the regulatory regions of several C-signal-dependent genes have been

characterized. Like these genes, the operon identified by Tn5 lac 94514 is expressed at

about the same time during development, but its expression does not depend on C-
signaling. The developmental regulation of this operon has been investigated. Based on
sequence similarity, the (24514 operon may encode an enzyme involved in glutamate
fermentation and a transcriptional regulator. The promoter of the operon shares similarity
with promoters that are expressed during growth and the 04514 promoter can be
transcribed in vitro by the major vegetative RNA polymerase. Developmental regulation
of £245 14 expression is complex, involving negative regulation by the product of the first
gene in the operon and positive regulation by one or more transcriptional activators.
Developmental expression of the (24403 gene is absolutely dependent on C-
signaling and on DNA sequences extending to —80 relative to the start site of
transcription. A search for regulatory proteins that interact with the upstream region has
been carried out. No protein likely to mediate C-signal-dependent expression of (24403

was identified.

To my husband, Enoch, my parents, and my son, Calvin, with love.

iv

ACKNOWLEDGEMENTS

In the many years that I endeavored to accomplish this goal, I have had a great
deal of support and encouragement from a lot of wonderful people.

I am especially indebted to my thesis advisor, Dr. Lee Kroos. It is a privilege for
me to have learned science and research under his guidance. I thank him for his support,
understanding and patience during these years. I also like to thank Mary Kroos for her
friendship.

I want to thank the members of my committee: Dr. Zachary Burton, Dr. Jon
Kaguni, Dr. Larry Snyder and Dr. Steven Trizenberg for their helpfiil advice and
suggestions regarding this dissertation project.

I also want to thank members of the Kroos lab. For the myxo team, I thank Makda
Fisseha for her kindness and fiiendship, Janine Brandner for being such a cheerful person
that makes working in the lab fun, Dvora Biran for carrying out the in vitro transcription
experiments and Gregory Velicer for his fiiendship and for constructing the orfl mutant
strain. I thank Nicco Yu, Bin Zhang, Hiroshi Ichikawa and Michelle Anderson (the
bacillus team) for always being resourceful of sharing ideas and reagents and for their
friendship.

My stay at East Lansing would not have been the same without the fiiends I have
made here. I would like to thank brothers and sisters fi'om the Lansing Chinese Christian
Ministry, especially, Anne Tseng, Shu-chin Lee, Shu-shin Chou, for their prayers,
support, care and fiiendship.

I could not have finished my thesis writing without the help from fiiends at

Columbus Chinese Christian Church. Numerous people have helped me to baby-sit

Calvin so that I could have time to write. I thank Dennis and Timmy Ong, Julie Wu, Katy
Yang and Fom-J em Lee for their loving hearts and continuous prayers.

My family has always been a source of support behind me. I thank my parents for
their unconditional love, support and encouragement. I thank my husband, Enoch, for his
love, fiiendship, encouragement, and patience over the years. Without his support and
understanding, I could not have reached this point. I am looking forward to the time

together with him and our son Calvin, who is truly a blessing fiom above.

TABLE OF CONTENTS

LIST OF TABLES .......................................................................... x
LIST OF FIGURES ......................................................................... xi
LIST OF ABBREVIATIONS ............................................................. xiii
INTRODUCTION .......................................................................... 1

CHAPTER I. Literature Review

Motility .............................................................................. 4
Fruiting body formation and sporulation ....................................... 6
A-signaling .......................................................................... 9
B-signaling .......................................................................... 16
D-signaling .......................................................................... 16
E—signaling .......................................................................... 17
C-signaling .......................................................................... 19
Regulation of gene expression ................................................... 27

CHAPTER II. Characterization of regulatory mechanism of a cell interaction-dependent
gene in Myxococcus xanthus

Abstract .............................................................................. 34
Introduction ......................................................................... 35
Materials and Methods ............................................................ 39
Results ............................................................................... 53
Localization of the developmental promoter controlling
Q4514 expression ......................................................... 53
Localization of an mRNA 5' end upstream of 945 14 ............... 59

vii

DNA sequence of the 04514 upstream region ........................ 62
Precise localization of the W4514 mRNA 5' end ..................... 66
Further deletion analysis of the Q4514 upstream region ............ 67

(24514 can be transcribed by RNA polymerase isolated fiom
growing M. xanthus cells ................................................ 72

Effect of an insertional mutation in ORFl on 04514 expression... 77

Production of ORFI in E. coli and testing for binding to the 04514
promoter .................................................................... 80

ORFl protein levels during M. xanthus growth and development. 83
Sporulation efficiency of Q45 14 mutants .............................. 86
Discussion ........................................................................... 92

CHAPTER III. Identification of regulatory proteins involved in the expression of a C-
signal-dependent gene

Abstract .............................................................................. 105
Introduction ......................................................................... 106
Materials and Methods ............................................................ 109
Results .............................................................................. 114
Further deletion analysis of the Q4403 promoter region ............ 114
Detection of a development-specific shified complex with the
04403 promoter region ................................................... 114
Competition assays with unlabeled fragments ........................ 117
Developmental timing of formation of the shifted complex ......... 117

Dependence of the DNA-binding activity on extracellular signals. 1 17

Localization of DNA sequence in the W4403 promoter region
required for binding ...................................................... 124

viii

Binding to other promoters .............................................. 129

Discussion ........................................................................... 132
SUMMARY AND PERSPECTIVES .................................................... 137
BIBLIOGRAPHY .......................................................................... 141

ix

LIST OF TABLES

Table 2.1. Bacterial Strains and Plasmids ................................................ 40

Table 3.1. Bacterial Strains and Plasmids ................................................ 110

LIST OF FIGURES

Figure 1.1. Dependence of developmental markers on A, B, C, and D factors... 11
Figure 1.2. Model depicting cellular responses to C-signaling ...................... 24

Figure 2.1. Physical map of the Q4514 insertion region and summary of
deletions tested for promoter activity ...................................... 55

Figure 2.2. Developmental expression of lacZ under the control of the Q4514
Promoter ....................................................................... 58

Figure 2.3. Localization of an mRNA 5' end within the Q4514 upstream region. 61

Figure 2.4. Nucleotide sequence of 1.9-kb of DNA upstream of Tn5 lac Q4514. 64

Figure 2.5. Primer extension analysis of Q45 14 mRNA ............................. 69
Figure 2.6. Developmental expression of lacZ from a small segment containing

the Q4514 promoter .......................................................... 71
Figure 2.7. In vitro transcription from the Q4514 promoter ......................... 74
Figure 2.8. Western blot analysis of cAin growing and developing M. xanthus

Cells ............................................................................ 76
Figure 2.9. Effect of orfl and 0112 mutations on developmental lacZ expression

under the control of the Q4514 promoter ................................ 79
Figure 2.10. Production of ORF1-6xHis in E. coli and affinity purification ...... 82
Figure 2.11. Detection of ORFl protein in extracts of growing M. xanthus ...... 85

Figure 2.12. Level of ORFl protein and B-galactosidase specific activity

during development of DK45 14 .......................................... 88
Figure 2.13. Sporulation of Q45 14 mutants ........................................... 91
Figure 2.14. Comparison of promoter sequences ..................................... 97
Figure 2.15. A model describing regulation of the Q4514 operon .................. 102

Figure 3.1. Developmental expression of lacZ from a DNA segment containing

Figure 3.2.
Figure 3.3.
Figure 3.4.
Figure 3.5.

Figure 3.6.

Figure 3.7.

the Q4403 promoter ......................................................... 116

Gel mobility shift analysis of the Q4403 promoter region ............. 119
Competition with unlabeled DNA .................. . ..................... 121
Developmental timing of formation of the shifted complex ........... 123
Dependence of the DNA-binding activity on extracellular signals... 126
Gel mobility shift assays of 5' deletion fragments containing the

Q4403 promoter .............................................................. 128
Gel shift experiments with different promoters .......................... 131

xii

dGTP
DNA
DTT
ECF
EDTA
HEPES
IPTG

kb

LB
mRNA
NAD+
NADH
ONPG
ORF
PAGE

PCR

LIST OF ABBREVIATIONS

ampicillin

adenosine-5'-triphosphate

base pair

coenzyme A

deoxyguanosine—S’-triphosphate

deoxyribonucleic acid

dithiothreitol

extracytoplasmic function
(ethylenedinitriol)tetraacetic acid
N-(2-hydroxyethyl)piperazine-N'-(2-ethanesulfonic acid)
isopropyl B-D thiogalactopyranoside

kilobases

kilodalton

kanamycin

Luria-Bertan:

messenger ribonucleic acid

nicotinamide adenine dinucleotide (oxidized form)
nicotinamide adenine dinucleotide (reduced form)
o-nitrophenol-B-D-galactoside

open reading frame

polyacrylamide gel electrophoresis

polymerase chain reaction

xiii

(p)ppGpp
PMSF

RNAP
RNase
SDS
Tc
TDT
Tris

tRNA

guanosine tetra- or penta-phosphate
phenylmethylsulfonyl fluoride

RNA polymerase

ribonuclease

sodium dodecylsulfate

tetracycline

terminal deoxynucleotide transferase
tris(hydroxymethyl)aminomethane

transfer ribonucleic acid

xiv

INTRODUCTION

The gram-negative soil bacterium Myxococcus xanthus displays remarkable social
behavior upon nutrient depletion, as cells move coordinately to aggregation centers and
form fruiting bodies. Inside each fruiting body, some of the rod-shaped cells lyse,
whereas others differentiate into ovoid-shaped spores. This morphological change is
accompanied by a highly ordered program of developmental gene expression. At least
five extracellular signaling interactions regulate the developmental process. To begin to
understand how cell-cell signaling interactions regulate developmental gene expression,
the regulatory mechanisms of two developmental genes have been investigated.

Chapter one presents a review of cell-cell interactions involved in Myxococcus
xanthus growth and development, as well as transcriptional regulation in M. xanthus.

Chapter two describes the investigation of the regulatory mechanism of a C-
signal-independent gene identified by Tn5 lac Q4514. Q4514 is expressed during
development with similar timing as several C-signal-dependent genes. However, its
promoter resembles vegetative promoters in sequence and can be transcribed by the
major vegetative M. xanthus RNA polymerase in vitro. Regulation of the 04514 operon
is complex, involving negative autoregulation by one of its own gene products and
positive regulation by one or more transcriptional activators. The content of this chapter
will be submitted to the Journal of Bacteriology with Dvora Biran and Gregory Velicer
as second and third authors, respectively. Dvora Biran was responsible for the in vitro

transcription experiments. Gregory Velicer constructed the orfl disruption strain and

tested developmental activity from the small Q4514 promoter in 0rf1 mutants. He also
constructed strain MGVO3 and tested it for promoter activity.
Chapter three describes the search for a regulatory protein(s) that mediates

developmental expression of Tn5 lac (24403, a C-signal-dependent gene. DNA between

—80 and —72 is crucial for C-signal—dependent expression of (24403. No protein that
interacts specifically with this region was identified.

The Summary and Perspectives section contains the conclusions and significance
of these results, and proposals for fiiture research directions. Both 04514 and (24403
require upstream regulatory elements for expression; however, no similar sequence has
been identified that is shared between the two genes. It is very likely that different
transcriptional activator proteins increase expression of these genes in response to

developmental signals.

Chapter I

Literature Review

Cell-cell interactions play a critical role in cellular differentiation in multicellular
organisms. Myxococcus xanthus, a gram-negative rod-shaped soil bacterium, exhibits
multiple forms of cell interactions by feeding, moving and developing cooperatively.
Some of the signals have been identified chemically, other signaling interactions can be
inferred from the behavior of cells. Cell-cell interactions are important for M. xanthus
cells to search for nutrients by moving in large groups known as swarms. Growth of M.
xanthus cells is also cell interaction-dependent, showing density dependency, which is
attributed to the increased concentration of nutrients available for growth that results
from the increased concentration of extracellular hydrolytic enzymes secreted by cells at
high cell density (Rosenberg et al., 1977). Cell-cell interactions are also manifested
during development when multicellular structures known as fruiting bodies are formed.
Since M. xanthus is genetically simple and readily amenable to genetic and biochemical
approaches, it provides a good experimental system for investigating how cell-cell
interactions control cellular differentiation during multicellular events, such as
development.

Motility

Myxococcus xanthus cells glide on a variety of different surfaces, forming swarms
that secrete lytic enzymes to digest prey bacteria. Within the swarm, movement of a small
group of cells is known as social motility (S). Occasionally, individual cells briefly move
outside of a swarm before returning, and this is known as adventurous (A) motility. Cell
motility is also indispensable for multicellular development. Nonmotile mutants are
unable to aggregate and form fruiting bodies and spores (Kroos et al., 1988; McBride,

1993). At least five classes of mutations that affect gliding motility have been identified

and characterized. These mutations have been grouped into five gene classes: A
(adventurous gliding), S (social gliding), mgl (mutual gliding), frz and dif (chemotaxis).

A mutation in any of the A genes results in cells that are able to move in groups,
but cannot glide as individuals. Conversely, a mutation in any of the S genes results in
cells that are able to move only as individuals (Hodgkin and Kaiser, 1979a). The A and S
systems work additively (i.e., strains with defects in both systems (A'S') are nonmotile).
There are at least 37 loci required for A motility (Hodgkin and Kaiser, 1979a; MacNeil et
al., 1994). One subset of A genes likely encodes proteins that are localized to the cell
envelope or extracellular matrix (Kalos and Zissler, 1990; Rodriguez and Spormann,
1999). The S system includes a minimum of 19 loci (Hodgkin and Kaiser, 1979b;
MacNeil et al., 1994) and S motility is operative only when cells are in close proximity to
each other (Kaiser, 1979; Kaiser and Crosby, 1983). S motility is correlated with pili,
which are long, thin fibers found in tufts at one pole of the cell. A motility, on the other
hand, is unrelated to pili. Components of the S system appear to be homologs of genes
involved in the biosynthesis, assembly and function of type IV pili in Pseudomonas
aeruginosa (Wu and Kaiser, 1995; Wu and Kaiser, 1997a).

Both A and S motility is abolished by a single mutation in the mgl locus, which
contains two genes, mglA and mng (Stephens and Kaiser, 1987; Stephens et al., 1988).
Mutations in mglA render cells completely nonmotile. It appears that MglA protein does
not function as a component of the gliding motor (Hartzell and Kaiser, 1991a), nor does it
regulate the transcription of A and S genes (MacNeil et al., 1994). Rather, MglA shares
amino acid similarity to GTP-binding proteins, suggesting a possible role of MglA in

signal transduction involved in motility (Hartzell and Kaiser, 1991b). Mutations in mng

result in cells with substantially reduced motility and reduced levels of the MglA protein.
The mng gene is co-transcribed with mglA and a normal amount of mgIBA transcript is
produced in mng mutants (Hartzell and Kaiser, 1991b). Because the transcription of
mglA is not affected by mng mutations, the reduced level of MglA could be due to
reduced stability of the MglA protein in the absence of Mng. Part of the predicted amino
acid sequence of Mng shows similarity to a calcium-binding site of yeast calmodulin,

but the function of Mng is still unclear (Hartzell and Kaiser, 1991b).

Bacterial chemotaxis-like sensory transduction systems control reversal of
gliding motility in M. xanthus. Two sensory processing systems, fiz and dif, homologous
to the Che chemotaxis system of E. coli, have been identified by sequence comparison
and biochemical characterization (Shi and Zusman, 1995; Yang et al., 1998). The frz
genes appear to regulate gliding movement of both individual cells and cells in groups.
The dif genes, in contrast, affect only S system-dependent gliding.

Fruiting body formation and sporulation

Myxococcus xanthus cells undergo a multicellular developmental process in
response to nutrient limitation (Shimkets, 1990). Within 4 to 8 hours after the onset of
starvation, the cells begin to aggregate by gliding to form mound-shaped structures
known as fruiting bodies. About 105 cells participate in the formation of a single fruiting
body. Within a fruiting body, some of the cells lyse, whereas by 24 hours, a small
population of motile, rod —shaped cells differentiate into non-motile, spherical spores that
are resistant to harsh conditions such as heat, freezing, desiccation and UV irradiation.
Aggregation and sporulation are temporally separated and sporulation in wild-type cells

occurs only after cell movements have led to the formation of fruiting bodies. The earliest

stages of aggregation are often accompanied by a form of organized cell movement
called rippling. During rippling, a series of equally spaced ridges of cells move in a
coordinated fashion like traveling waves (Shimkets and Kaiser, 1982). Rippling is not
absolutely required for the construction of fi'uiting bodies or sporulation, but normally
accompanies fruiting body formation (Shimkets, 1990).

Multicellular development in Myxococcus xanthus depends on at least five
extracellular signals known as the A-, B-, C-, D-, and E-signals. Signaling mutants were
identified by isolating conditional nonsporulating mutants, whose sporulation can be
rescued by codevelopment with wild-type cells or mutants of a different signaling group
(Hagen et al., 1978). The signaling mutants are defective in producing a signal, but not in
perceiving that signal. Mutants unable to produce any one of the five signals are arrested
in development at a particular stage. By examining the expression of developmental
reporters in signaling defective mutants, it has been shown that the A- and B-signals are
required at the onset of development, the D- and E-signals are required at 3 to 5 hours
after the onset of development, and C-signaling is required at about 6 hours into
development (Downard et al., 1993; Kaiser and Kroos, 1993; Downard and Toal, 1995).
Of the five signals, only the A-signal has been defined biochemically.

Fruiting body morphogenesis and sporulation are accompanied by the coordinated
expression of many developmentally regulated genes and the synthesis of new proteins.
More than 30 proteins, identified as bands separated by gel electrophoresis, as well as
several new cell surface antigens are associated with developing cells (Inouye et al.,
1979b; Gill and Dworkin, 1986; Gill et al., 1987). Among the most abundant proteins

expressed during development are proteins S (a spore coat protein) and 8-1 (a protein

found inside spores) (Inouye et al., 1979a; Teintze et al., 1985). These two proteins are
encoded by tps and ops, respectively, and these genes are located next to each other,
separated by a 1.4 kb DNA segment. Proteins S and 8-1 are 90% identical, but they are
produced at different times during development (Inouye et al., 1983a; Inouye et al.,
1983b). At 5 hours after starvation, tps begins to be expressed and later the protein S
produced self-assembles the outermost spore coat. The ops gene is not expressed until 24
hours into deve10pment (Downard et al., 1984; Downard and Zusman, 1985). Another
protein produced during development is myxobacterial hemagglutinin, a protein with
lectin activity, which appears at 10 hours (Romeo and Zusman, 1987).

Many developmentally regulated genes have been identified by genomic insertion
of Tn5 lac, a transposon containing a promoterless lacZ that generates transcriptional
fusions to promoters in the chromosome (Kroos and Kaiser, 1984). Among 2374 Tn5 lac
insertions, 29 insertions showed increased B-galactosidase activity upon starvation,
implying that about 1% of the genome is up-regulated during fruiting body development
(Kroos et al., 1986). Only a small portion of the 29 Tn5 lac insertions showed
developmental defects that impair aggregation or sporulation (Kroos et al., 1990). A
similar low frequency has been observed in the eukaryotic slime mold Dictyostelium
discoideum, which has a multicellular development like that of Myxococcus xanthus
(Kuspa and Loomis, 1992). Possible explanations are that subtle changes in morphology
might escape notice and/or that many genes are at least partially redundant in their
functions. B-galactosidase activities from the 29 developmentally regulated fusions have
been assayed in wild-type cells as well as in cells that are defective in an extracellular

signaling interaction (Kroos and Kaiser, 1987; Downard et al., 1993; Kaiser and Kroos,

1993; Downard and Toal, 1995). Dependency of each fusion on extracellular signaling is
depicted in Figure 1.1. Expression of most fiisions is dependent upon A- and B- signaling.
Fusions expressed around 4 hours require D-signaling and nearly all genes that begin to
be expressed after 6 hours into development appear to be at least partially C-dependent.
Expression of fps, which normally begins around 5 hours, is reduced in E—signaling
mutant cells, indicating that the E-signaling interaction affects the pattern of gene
expression by 5 hours into development (Downard et al., 1993).
A-signaling

A-signaling mutants are arrested early in development, prior to aggregation. The
A-signal itself has been purified from media conditioned by developing cells by
monitoring the expression of Tn5 lac fusion Q4521, whose expression requires A-
signaling and therefore could serve as an assay to identify fractions containing A-signal
activity. The A-signal is composed of a subset of amino acids that can fimction either
individually or in combination at a concentration greater than 10 M (Kuspa et al., 1992).
Fifteen amino acids have A-factor activity. The six amino acids with the highest specific
activities are tyrosine, proline, tryptophan, phenylalanine, leucine and isoleucine.
Peptides also have activity (Plamann et al., 1992). Peptides most likely generate A signal
as they are degraded to their constituent amino acids by extracellular and periplasmic
proteinases, because the activity of each peptide tested is approximately equal to the sum
of the activities of its amino acids. It appears that proteinases, released early during
development, are involved in generating the A-signal by degrading surface proteins to

their constituent amino acids and peptides. All of the proteinases tested have the ability to

Figure 1.1 Dependence of developmental markers on A, B, C, and D factors. Each
marker is placed above an arrow positioned along the x axis according to the time at
which its expression normally begins during development. Position along the y axis
indicates the marker’s dependence on cell-cell interactions, as determined by measuring
its expression in asg, bsg, csg and dsg mutants. For example, a marker whose expression
is reduced in an asg mutant exhibits partial dependence on A factor (abbreviated “partial
A”). Similarly, a marker whose expression is abolished in a bsg mutant exhibits absolute
dependence on B factor (abbreviated “absolute B”). Dependencies are cumulative, so a
higher position along the y axis indicates greater dependence on cell-cell interactions.
Tn5 lac insertions are indicated by 0 followed by a four digit number, and brackets
connect insertions that probably lie in the same transcription unit. Reprinted with
permission from authors. Kaiser, D. and Kroos, L. (1993) In Intercellular signaling.

(Dworkin, M. and Kaiser, D., eds) 257-283, American Society for Microbiology.

10

04 506 04459 hydrophme

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

shttt
ops
ABSOLUTE D 04401
04497
E04435
04500 94480
04406 ' l:04403 04529
ABSOLUTE c '
04531
04499 E04473
04400 04414 04474
ABSOLUTE B
_ mbhA
ABSOLUTE D
alkaline
04427 04514 04492 phosphatase
PARTIAL c
Ips
PARTIAL oi
04494
04491
04427 04530
04457 04411 neutral
04:21 04445 phosphatase
ABSOLUTE A PARTIAL D
6394 PARTIAL A
PARTIAL c
04469
PARTIAL o
csgA
04455
04406
PARTIAL B
protein C
ABSOLUTE A
PARTIAL D
acid
phosphatase
PARTIAL A
dFA-I
l L I I l 1 :H—l
o 2 4 6 6 10 12 14 16 16 25 27

HOURS OF DEVELOPMENT

1]

generate A-signal activity and at least two different proteinases can be isolated from
developing cell extracts (Plamann et al., 1992).

The generation of extracellular A-signal requires the products of at least five asg
genes. These include asgA, asgB, asgC, and two newly characterized genes, asgD and
ang (Kuspa and Kaiser, 1989; Cho and Zusman, 1999; Garza et al., 2000). A mutation
in any of these genes renders cells defective in generating A—signal, but the cells maintain
their ability to respond to extracellular A-signal. The asgA gene encodes a histidine
protein kinase that resembles the type found in sensor proteins of two-component signal
transduction systems (Plamann et al., 1995). The AsgA protein is unusual in that it also
contains a receiver domain typically found in the response regulator protein of two-
component systems. It has been proposed that AsgA may fiinction in a multicomponent
phosphorelay-type signal transduction mechanism that senses starvation and responds by
altering the expression of genes required for production of A-signal (Plamann et al.,
1995)

The asgB gene encodes a putative DNA-binding protein with a helix-tum-helix
(HTH) motif near its C-terminus (Plamann et al., 1994). The predicted HTH motif in
AsgB is highly similar to the HTH found in region 4.2 of major sigma factors, which
recognizes and interacts with the -35 region of promoters. It has been proposed that AsgB
may act as a transcriptional activator or repressor (Plamann et al., 1994).

The asgC gene encodes the major sigma factor of RNA polymerase holoenzyme,
oA (Davis et al., 1995). The mutant allele, asgC 76 7, results in a glutamate to lysine
substitution in region 3.1. This region is suspected to be involved in (p)ppGpp-mediated

gene regulation in E. coli (Harris et al., 1998). One possible explanation for the A-

12

signaling defect of asgC 7 6 7 cells is that the mutated oA fails to respond to the changing
levels of (p)ppGpp and the expression of (p)ppGpp-dependent genes is impaired.

The AsgD protein, like AsgA, is predicted to be a component of a signal
transduction system because it has a receiver domain located at its N-terminus and a
histidine protein kinase at its C-terminus (Cho and Zusman, 1999). Fruiting body
formation and sporulation are arrested when asgD mutant cells are plated on CF medium,
a low-nutrient medium that induces development upon starvation. However, when the
mutant is plated on stringent starvation medium rather than CF, cells are able to form
fruiting bodies, suggesting that AsgD is directly or indirectly involved in sensing
nutritionally limiting conditions.

Another gene involved in generating A-factor is ang (Garza et al., 2000). A-
factor consists of a mixture of amino acids, which is generated by extracellular proteases
that degrade cell surface proteins into constituent amino acids. The ang mutant yields
about lO-fold less extracellular protease activity and about twofold less A-factor than
wild-type cells during development (Garza et al., 2000). These results suggest that the
primary defect of ang cells is in the production or release of extracellular proteases.

Insight into how cells respond to A-signaling was obtained by studying mutants

that result in constitutive expression of the A-signal-dependent reporter gene, £24521, in

 

i F'i. .

asg' cells. 94521 was identified as a developmentally regulated Tn5 lac insertion and its
expression is abolished in any of the A-signal mutants. Genetic suppressors that allow A-
signal-independent expression of (24521 in the asgB mutant background have been
identified. Three loci have been characterized so far, known as sasA, sasN, and sasS. The

sasA locus is required for biosynthesis of the O-antigen of M. xanthus lipopolysaccharide

13

(Guo et al., 1995). The connection between the absence of O-antigen and A-signal-
independent expression of 94521 is unclear. The sasN locus encodes a novel protein with
no homologue in the databases (Xu et al., 1998). The N-terminus of SasN appears to
contain a highly hydrophobic region and a leucine zipper motif. The sasN-null mutant
expressed 04521 at a higher level during both growth and development, as compared
with wild-type cells, suggesting that SasN negatively regulates Q4521 expression during
growth and development. The sasS locus appears to encode a sensor histidine protein
kinase of a two-component system (Yang and Kaplan, 1997). The C-terminus of SasS
contains all of the conserved residues found in sensor histidine protein kinases, and the
N-terminus of SasS contains two putative transmembrane domains that might localize
this protein to the cytoplasmic membrane. The sasS-null mutants are defective in fruiting
body formation and sporulation and express Q4521 at a basal level. These data suggest
that SasS may sense extracellular A-signal and positively regulate Q4521 expression and
expression of other genes induced during M. xanthus development. Two other putative
positive regulators of A-signaling responses, SasR, an NtrC-like response regulator, and
SasP, have been identified by a separate approach (Guo et al., 2000). Disruption in sasR
or sasP abolished Q4521 expression during development in the presence or absence of
lipopolysaccharide O-antigen.

Q4521 expression depends on cell density during starvation (Kuspa et al., 1992).
At cell densities below 2x108 cells/ml, Q4521 is expressed at 25% or less of the
maximum level. The expression of £24521 increases drastically from about 5x108
cells/ml, reaching its maximum level at about 109 cells/ml. The low expression at low cell

density appears to be due to less A-signal activity since Q4521 expression is restored to

14

the maximum level when A—signal is added to cells at low cell density. These results
suggest that the A-signal level is proportionate to the cell density and that A-signaling
monitors whether a sufficient density of starved cells is present to make a fruiting body.
To arrest growth and progress into development, M. xanthus cells must sense
nutrient limitation as well as high cell density. In E. coli, when nutrient is limited,
uncharged tRNA binds to the accepter site of ribosomes and the highly phosphorylated
guanosine tetra- or penta-phosphate, (p)ppGpp, is formed (Cashel et al., 1996). Because
(p)ppGpp is made by ribosomes when they stall due to the lack of any amino acid,
generation of ppGpp is rich with nutritional information. A similar scenario is seen
during M. xanthus development. Within 30 minutes of starvation, the level of (p)ppGpp is
elevated in response to amino acid limitation (Singer and Kaiser, 1995). Singer et al.
expressed the E. coli relA gene encoding (p)ppGpp synthetase I inM xanthus and found
that development-specific lacZ reporters are induced without starvation (Singer and
Kaiser, 1995). This result suggests that (p)ppGpp alone is sufficient to initiate
developmental gene expression. A mutant that lost the ability to accumulate (p)ppGpp in
response to starvation failed to form fruiting bodies and spores (Manoil and Kaiser,
1980), and these defects were rescued by providing (p)ppGpp (Harris et al., 1998). The
M. xanthus relA homolog has been cloned and this gene rescued the developmental defect
of the above mutant (Harris et al., 1998). Disruption of the M. xanthus relA gene rendered
cells unable to accumulate (p)ppGpp and prevented fruiting body development. These
results support the idea that elevated (p)ppGpp is sufficient to initiate the developmental
program. Harris et al. also showed that the accumulation of (p)ppGpp was necessary and

sufficient to induce A-signal production (Harris et al., 1998). A-signal is not produced in

15

the M xanthus relA -null mutant that fails to accumulate (p)ppGpp, and expression of the
E. coli relA gene in M. xanthus restores production of A-signal. It has been proposed that
nutrient limitation results in (p)ppGpp production, which triggers A-signal production,
and the level of A-signal serves as a cell density indicator to determine whether sufficient
cells are present to undergo development.
B-signaling

Another signaling interaction required early in development is B-signaling.
Expression of nearly all of the developmental markers examined is either reduced or
abolished by B-signaling mutations, suggesting that B signal acts early, within 2 hours
after the onset of development. B-signaling mutants fail to form fi'uiting bodies and
spores (Gill and Bornemann, 1988). A subset of the B-signaling mutations map to the
bsgA gene, which encodes an ATP-dependent protease with a high degree of amino acid
similarity to protease La (Lon) of E. coli and Bacillus brevis (Gill et al., 1993). BsgA is a
cytoplasmic protein. Its role in generating the B-signal and the mechanism of
extracellular complementation of B-signaling mutants are unknown. It has been proposed
that the defect in intracellular proteolysis in bsgA mutants is involved in generating the B-
signal, but the chemical nature of the B-signal is unknown. Proteolysis is involved in the
activation of some regulatory proteins, such as the development-specific sigma factors 0E
and UK of B. subtilis (Kroos and Cutting, 1994). BsgA may affect a regulatory protein
and/or the B-signal itself through proteolysis.
D-signaling

D-signaling mutants arrest development around the stage of aggregation. Under

nutrient limitation, the mutants form larger, less compact aggregates than wild-type cells

16

and sporulation is delayed and substantially reduced. The mutants are defective in B-
galactosidase expression from Tn5 lac fusions whose expression normally begins about 5
to 6 hours into development (Cheng and Kaiser, 1989b). Two dsg point mutations map to
the same locus, dsgA (Cheng and Kaiser, 1989b). The amino acid sequence of DsgA has
50% identity to the sequence of translation initiation factor IF3 of E. coli and Bacillus
stearothermophilus, a protein which enables the ribosome to select the initiation codon
(Cheng et al., 1994). It has also been shown that DsgA firnctions like translation initiation
factor IF3 of E. coli in M. xanthus (Kalman et al., 1994). Although cells with dsgA point
mutations are defective in development, they grow at the same rate as wild-type cells.
Complete disruption of the dsgA gene by Tn5 insertion is lethal (Cheng and Kaiser,
1989a). This suggests that the modified proteins encoded by the dsgA point mutants
retain the functions of IF3 required for growth. The developmental defect of these
mutants is not due to less DsgA protein during development since the level of DsgA is
uniform throughout growth and development (Kalman et al., 1994). Rather, the
developmental defect must result from the altered activity of the mutant DsgA proteins.

The dsgA gene product is probably not the signal itself. Rather, as part of the
translational machinery, it may be involved in generating the signal. Rosenbluh and
Rosenberg have shown that a mixture of fatty acids rescues the fruiting body formation
and sporulation of clsgA mutants (Rosenbluh and Rosenberg, 1989), suggesting a possible
role of fatty acids in D-signaling. Alternatively, the rescue could be explained by the idea
that fatty acids alter the permeability of the cell membrane and allow the transfer of
another molecule(s), such as D-signal.

E-signaling

l7

Another class of signaling mutants was identified in the study of regulation of the
tps gene, which encodes spore protein S (Downard et al., 1993). The development of E-
signaling mutants is arrested at about 3 to 5 hours after the onset of starvation. The
mutants are defective in expression of fps and in aggregation, fi'uiting body formation and
sporulation. The defects in gene regulation and sporulation were rescued by mixing with
wild-type cells, but the defect in fi'uiting body formation was not corrected. Mutations
affecting E-signaling mapped to the esg locus. Two open reading frames have been
identified at this locus, which share sequence similarity with the Ela and Elb subunits of
a multienzyme complex, branched-chain keto acid dehydrogenase (BCKAD). BCKAD is
involved in the synthesis of short branched-chain fatty acids (BCFAs) from branched-
chain amino acids (BCAAs) (Toal et al., 1995). These BCKAD products are precursors in
the biosynthesis of long branched-chain fatty acids, which are incorporated into
phospholipids during vegetative growth. Functional analyses using an esg::Tn5 insertion
supported the results of sequence comparisons because the mutant had increased BCAAs
due to a lack of BCKAD activity. Furthermore, when the mutant cells were grown
vegetatively in the presence of short BCFAs, the defects in development were corrected,
suggesting that short BCFAs allowed cells to bypass the metabolic block caused by the
‘esg mutation. This result also suggests that BCFAs synthesized during vegetative growth
affect the developmental process later, either directly or by effecting production of other
molecules. Since a null mutation in esg reduced but did not abolish the production of
short BCFAs, an alternative pathway to generate these compounds must also exist.

What is the nature of E-signal? A model has been proposed that the BCFAs,

synthesized during growth, are released from cellular phospholipid by a developmentally

18

regulated phospholipase during fruiting body formation (Downard and Toal, 1995), and
that one or more of the released BCFAs constitutes E-signal. Alternatively, fatty acids
may not serve as the signal directly. Rather, they may act through modifying other
molecules, such as proteins, to participate in generating the E-signal.
C-signaling

Among the five identified signaling interactions, C-signaling affects development
at the latest stage based on the developmental phenotype and the effect on developmental
gene expression. Upon starvation, C-signaling mutant cells are arrested early in
aggregation. C-signaling mutants fail to form ripples, tight mounds, or spores. Rather,
they form diffuse and flat mounds around 18 hours into development, while tight mounds
are formed around 12 h by wild-type cells. Sporulation is severely impaired in C-
signaling mutants and the expression of genes normally induced at six hours or later in
development is reduced or completely abolished (Hagen et al., 1978; Shimkets and
Kaiser, 1982; Kroos and Kaiser, 1987).

All of the C-signaling mutations map to a single gene, csgA (Shimkets and Asher,
1988). The csgA gene encodes a 25-kD protein with sequence similarity to the members
of the short chain alcohol dehydrogenase (SCAD) family (Lee et al., 1995). These
enzymes use NAD(H) or NADP(H) to catalyze the interconversion of secondary alcohols
and ketones or to mediate decarboxylation. They have a wide variety of substrates and
are involved in the production of many different signaling molecules including steroids,
prostaglandins, and nodulation factors in plants (Persson et al., 1991; Baker, 1994). ngA
has a NAD(P) binding motif at its N-terminus. Disruption of this motif by amino acid

substitution abolished its ability to rescue csg' mutant cells (Lee et al., 1995). The activity

19

is also abolished by a substitution at the putative catalytic site of the enzyme (Lee et al.,
1995). Furthermore, addition of NAD(P) to a mixture of MalE-ngA and csgA mutant
cells increased sporulation of the csgA cells, while adding NAD(P)H to the mixture
reduce sporulation efficiency (Lee et al., 1995). These results suggest that enzymatic
activity of ngA is necessary for C-signaling. ngA appears to be unique among
members of the SCAD family in that it functions extracellularly. The substrate(s) for
ngA is unknown.

A 17 kD protein, named C-factor, has been purified from wild-type developing
cells that is able to rescue the developmental defects of csgA mutant cells after mixture
(Kim and Kaiser, l990c; Kim and Kaiser, 1990d). C-factor has been proposed to be
encoded by csgA based on the following findings. First, wild-type cells produce C-factor,
but csgA mutant cells do not. Secondly, a stretch of amino acid sequence in the middle of
C-factor protein was obtained and it corresponds exactly with a deduced amino acid
sequence in the middle of ngA. Thirdly, polyclonal antibodies raised against a lacZ-
csgA fusion protein and purified by binding to the fusion protein react with the 17 kD C-
factor. These findings argue that C-factor is encoded by csgA. However, the relationship
between the 17 kD C-factor and the 25 kD ngA is unknown. One possibility is that C-
factor is a proteolytic product of ngA.

The aggregation, sporulation and developmental gene expression defects in csgA
mutant cells can be rescued by codevelopment with wild-type cells or by the addition of
purified C-factor (Hagen et al., 197 8; Kim and Kaiser, 1990c; Kim and Kaiser, l990d).
Anti-ngA antibodies inhibited both fruiting body morphogenesis and sporulation of

wild-type cells unless they were first neutralized with purified C-factor (Shimkets and

20

Rafiee, 1990). Furthermore, the antibodies also localized ngA at the cell surface and in
the extracelluar matrix. These results indicate that ngA is a cell-surface-associated
protein and nondiffusible.

Rescue of csgA cells by wild-type cells requires direct cell-cell contact and both
the donor and receptor cells have to be motile for effective C-signaling (Kroos et al.,
1988; Kim and Kaiser, 1990b; Kim and Kaiser, 19900). The extracellular
complementation does not occur when wild-type and csgA mutant cells are separated by a
membrane (Kim and Kaiser, 19900). Nonmotile mgl mutant cells exhibit characteristics
of the C-signaling mutant phenotype despite the finding that the mgl mutant produces
normal amounts of C-signal. Addition of purified C-factor to the nonmotile cells induces
sporulation without restoring motility and the aggregation defect 00m and Kaiser, 1991).
Manually aligning nonmotile cells by creating narrow grooves on an agar surface restored
C-signaling (Kim and Kaiser, 1990a). These observations suggested that motility is
required to establish the prOper cell alignment needed for efficient C-signal transmission
(Kim and Kaiser, 1990a). Specifically, the alignment of cells may be important to
promote end-to-end contacts between cells (Sager and Kaiser, 1994).

There is evidence that C-signal induces changes in the motility behavior of
individual M xanthus cells. Addition of C-factor to csgA cells causes increased gliding
speeds, increased duration of the mean gliding interval, and decreased stop frequency
(Jelsbak and Sogaard-Anderson, 1999). These changes in cell motility are thought to help
move cells into aggregation centers efficiently during fruiting body formation. C-signal-
induced stimulation of motility depends on the cytoplasmic Frz signal transduction

system. Elements of the Frz system have sequence homology to components of the signal

21

transduction system for chemotaxis in E. coli. F rz proteins control the frequency of
gliding reversal, orienting the direction of M. xantlrus cell movement in response to
chemical stimuli (McBride et al., 1992; Shi et al., 1993). FrzCD, one of the proteins in
the Frz system, is homologous to Tar, a methyl-accepting chemotaxis protein of E. coli
(Dworkin, 1996). Exposure of developing csgA mutant cells to purified C-factor
increases the ratio of methylated to non-methylated FrzCD protein (Sogaard-Anderson
and Kaiser, 1996). Methylation depends on the methyltransferase FrzF, which is a cheR
homolog. C-factor-induced methylation also depends on FruA, a DNA-binding response
regulator that is necessary in vivo for all known responses to C-factor.

csgA mutant cells respond differently to different levels of added C-factor. C-
factor at a lower concentration rescues aggregation and expression of a gene partially
dependent on C-signaling, whereas a higher concentration of C-factor is needed to rescue
sporulation and expression of a gene absolutely dependent on C-signaling (Kim and
Kaiser, 1991). These observations suggested that C-signaling induces and coordinates the
temporally separated morphogenetic processes of aggregation and sporulation during
fruiting body formation. The csgA gene has a long upstream sequence and full expression
of csgA depends on multiple factors including ngA itself (Kim and Kaiser, 1991).
Positive autoregulation of csgA leads to an increased level of ngA later in development.
Not only does csgA expression increase in response to C-signaling, the C-signaling
interaction between cells also increases due to close and ordered packing of cells in the
nascent fruiting body. Therefore, ngA appears to act as an extracellular developmental
timer to bring about the appropriate order of the developmental process, first aggregation,

then sporulation.

22

A model has been proposed for the C-signal transduction pathway based on the
results of genetic analyses of mutants deficient in C-signal-dependent activities (Figure
1.2). In this model, the signaling event is the interaction between C-signal on one cell end
and a hypothetical sensor on a second cell end. This interaction activates the C-signaling
pathway in the second cell. The first known component downstream of the signaling
event, fruA, is required for all four C-signal-dependent activities: rippling, aggregation,
sporulation and developmental gene expression (Ogawa et al., 1996; Sogaard-Andersen
et al., 1996; Ellehauge et al., 1998). FruA is a DNA-binding response regulator that is
expressed at 3 to 6 hours after the onset of development (Ellehauge et al., 1998). The
expression of fruA is independent of C-signal, but depends on the earlier signals A and E.
However, genetic experiments suggest that C-signaling controls FruA activity post-
translationally by phosphorylation. Downstream of fiuA, the pathway branches, with one
branch leading to the motility responses to C-signaling, i.e., rippling and aggregation.
The frz signal transduction system is required for this branch. The other branch leads to
sporulation. The products of the devRS locus are involved in this branch. Mutants with
defects in devRS ripple and aggregate normally, but have impaired sporulation. Five
genes are present at this locus, orfI, orj2, devT, devR and devS, none of which share
sequence similarity with known proteins in the databases (Kaiser, 1999). Expression of
the dev operon is first detected afier 6 hours of development and is under strong negative
autoregulation. devS is essential for this repression (Kaiser, 1999). Cytometric
experiments have shown that dev is expressed in a bimodal pattern among cells, i.e.,
some cells are fully on and some cells are fiilly off, unlike some other developmentally

regulated genes (Russo-Marie et al., 1993). Fluorescence experiments indicate that in a

23

 

ngA.

 

 

 

 

 

 

\ F V
rippling
Frz + aggregation
aFruA~P d I .
1 Act 1 av -> sporu atIon

Q4400

< ngA 04403
£24499

1 L J

 

Figure 1.2. Model depicting cellular responses to

C signaling.

24

 

nascent fruiting body, the oval cells undergoing development in the center of the fruiting
body express dev, whereas the rod-shaped peripheral cells do not express dev (Kaiser,
1999). These results suggest that dev expression behaves like a switch at the cellular level.
Based on the facts that ngA synthesis is similar in dev mutant cells, in fruA mutant cells,
and in wild-type cells, and that csgA autoregulates, a third branch leading to csgA
expression must also exist (Kaiser, 1999). A newly identified regulator Actl is involved

in this pathway (Gorski et al., 2000). The act] mutant cells produce substantially less
ngA than normal. They aggregate normally, but fail to sporulate and expression of the
dev locus is severely impaired in the mutant. These results suggest that the reduced
amount of ngA in the act] mutant is adequate for aggregation, but not for sporulation. It
has been proposed that Act] responds to C-signal reception by increasing the expression
of the csgA gene, creating a positive feedback loop that boosts the level of ngA for later
developmental events.

Suppressors of csgA (soc) that restore sporulation of csg mutant cells also bring
insight about the mechanism of C—signaling. Several suppressors have been identified.
Two transposon mutations in the socABC operon restore development to csgA null
mutants (Lee and Shimkets, 1996). Mixing of these socABC and csgA double mutants
with csg cells stimulates the development of the csgA cells, suggesting that the suppressor
cells are able to produce C-signal or a similar molecule via a csgA-independent pathway.
Three genes are encoded in this operon: socA, like csgA, encodes a protein similar to
members of the SCAD family; socB encodes a putative membrane anchoring protein; and
socC has no homolog in the databases. SocA protein shares 28% identity and 51%

similarity in amino acid sequence with ngA. Both of the transposon suppressor

25

mutations inactivate socC, leading to a 30- to 100- fold increase in socA transcription,
suggesting that socC may encode a negative autoregulator of the socABC operon. It has
been proposed that the mutations cause derepression of socA and the overproduced SocA
substitutes for ngA.

Another suppressor mutation is named socD. The socD gene encodes a histidine
protein kinase and the suppressor mutation restores sporulation to csgA cells under
normal developmental conditions at 32°C (Rhie and Shimkets, 1991; Shimkets, 1999).
This mutation also induces sporulation of otherwise wild-type cells and csgA cells at
15°C under conditions that normally support growth. These results suggest that SocD
might be involved in nutrient sensing.

Investigation of another suppressor mutation, socE, has revealed a new role of
ngA in growth and development. Disruption of socE by a transposon restores
development to csgA mutants (Crawford and Shimkets, 2000). SocE is a vegetatively
expressed, highly basic protein with no homolog in the databases. SocE is essential for
vegetative growth in csgA + cells since attempts to transfer the socE null allele to csgA+
cells failed. Results from experiments in which socE was placed under the control of a
light-inducible promoter have shown that socE depletion arrests growth and induces
sporulation under normal growth conditions in an otherwise wild-type background. SocE
depletion also curtails DNA and RNA synthesis, inhibits cell elongation, and induces
accumulation of the stringent nucleotides [p]ppGpp. Amino acid substitutions in the
ngA coenzyme-binding pocket or catalytic site eliminated growth arrest. A relA
mutation also eliminated growth arrest. Pseudorevertants selected for grth following

socE depletion have mutations in csgA or relA. These results suggest that SocE works in

26

opposition to ngA to regulate growth arrest through relA, i.e., SocE inhibits the stringent
response and ngA induces it. This is supported by the fact that csgA cells have a very
low [p]ppGpp level, whereas socE csgA cells have a high [p]ppGpp level throughout
development, as do wild-type cells. ngA shares sequence similarity with members of the
SCAD family. Although its enzymatic activity appears to be necessary for growth arrest
of SocE-depleted cells, extracellular C-signaling is unlikely to be involved under the
conditions tested because the cells are dispersed at low cell density in grth medium,
which prevents contact-dependent exchange of C-signal.
Regulation of gene expression

Regulation of gene expression in response to various environmental cues is
mediated by a variety of mechanisms in prokaryotes. Much of the regulation occurs at the
transcriptional level. Many bacteria possess multiple sigma factors that combine with
core RNA polymerase to produce holoenzymes that express different sets of genes in
response to a wide variety of signals. A well-studied example of this is the sporulation
process of B. subtilis (cf. Haldenwang, 1995), which involves a cascade of sigma factors,
activated at different stages in development and in two different compartments, to
regulate genes spatially and temporally. Multiple sigma factors have also been identified
in M xanthus. The major sigma factor in vegetatively growing cells, designated GA, was
identified by using the E. coli 070 gene as a hybridization probe (Inouye, 1990) and RNA
polymerase containing CA has been purified from vegetative cells (Biran and Kroos,
1997). CA has sequence similarity to 07° of E. coli and 0'43 of B. subtilis. Partially purified
oA RNA polymerase or holoenzyme reconstituted from GA and M xanthus core RNAP

transcribed both vegetatively-expressed M xanthus genes that were tested.

27

The promoters of one vegetative gene, pilA (Wu and Kaiser, 1997b), and three
developmental genes, mbhA (Romeo and Zusman, 1991), sdeK (Garza et al., 1998), and
Q4521 (Keseler and Kaiser, 1995) resemble in the —24 and —12 regions of promoters that
are recognized by 054 in other bacteria. Mutational analysis of the (24521 promoter region
has shown that DNA base pairs shared with the consensus a"4 promoter sequence are
essential for Q4521 promoter activity, suggesting that it is likely to be transcribed by 054
RNA polymerase in vivo (Keseler and Kaiser, 1995). An rpoN gene encoding a 0'54
homolog has been cloned from M xanthus and attempts to make a null mutant have
failed, suggesting that the gene is essential for M xanthus growth, unlike in some other
organisms where 054 is nonessential (Keseler and Kaiser, 1997).

CarQ, a sigma factor of the extracytoplasmic fiInction (ECF) subfamily, is
responsible for light-induced synthesis of membrane-associated carotenoids inM
xanthus through the activation of its own carQRS promoter and the carC promoter. CarC
encodes for phytoene dehydrogenase that converts colorless phytoene into colored
carotenoids (Hodgson and Murillo, 1993; Gorham et al., 1996). The ECF sigma factors
are a subfamily of 070 factors that typically regulate genes encoding proteins with
extracytoplasmic functions (Lonetto et al., 1994). These sigma factors are shorter than
other known sigma factors and show significant divergence from other 070 family
members, most noticeably in regions 2.4 and 3. The activity of CarQ is negatively
regulated by an inner membrane protein, CarR, which is also encoded in the carQRS
operon (Gorham et al., 1996). CarR is unstable in the light and is essential for the

inactivation of the carQRS promoter in the dark. It has been proposed that the membrane-

28

associated CarR sequesters CarQ to the membrane in the dark to prevent it from
fiinctioning, whereas in the light, loss of CarR leads to release of the sigma factor.

Four other putative M xanthus sigma factors have been described, but no
promoters have been identified that are recognized by these sigma factors. SigB, a sigma
factor expressed later in sporulation, may be responsible for the expression of late
developmental genes (Apelian and Inouye, 1990). Expression of one such gene, ops,
which is expressed around 24 hours into development, was blocked in a sigB deletion
strain. However, no evidence of direct transcriptional regulation by SigB has been
described. sigB is not essential for development since sigB mutant cells exhibit normal
fruiting body formation and sporulation. However, sigB mutant spores show severe
defects in stability and viability, suggesting a role for SigB in maturation of myxospores.
SigC, another developmental sigma factor, is expressed immediately after cells enter
development and its expression is reduced significantly later, at the onset of sporulation
(Apelian and Inouye, 1993). sigC mutant cells develop normally, but sigC cells are
capable of forming fi'uiting bodies and myxospores on semi-rich media. This result
suggests that sigC may be involved in negatively regulating the initiation of fruiting body
formation. SigD is a sigma factor expressed during the stationary phase of vegetative
growth and early in development (U eki and Inouye, 1998). sigD mutant cells exhibit
growth defects during the late-log phase and stationary phase as well as an altered pattern
of protein synthesis. The deletion mutant also showed a significant delay in fruiting body
formation and sporulation and yielded fewer spores than wild-type cells. Additionally,

another sigma factor of the ECF subfamily, RpoEl, has been identified that may play a

29

role in transcriptional regulation of genes involved in motility behavior during both
vegetative growth and development (Ward et al., 1998).

Transcriptional activator proteins are involved in transcriptional regulation of
many genes in M xanthus. In all systems investigated, 054 promoters require the binding
of an activator protein to an upstream sequence. Likewise, the M xanthus 94521 and
mbhA genes require DNA upstream of the core promoter elements for developmental
expression (Romeo and Zusman, 1991; Keseler and Kaiser, 1995). Despite strong
similarity in the core promoter regions, expression of these two promoters differs
significantly in terms of the timing of expression during development and the
requirements for a solid surface and extracellular signals, suggesting the involvement of
different activator proteins.

Another example of transcriptional regulation by activator proteins comes from
the study of tps and ops gene expression (Kil et al., 1990). These two genes are separated
by 1.4 kb on the M xanthus chromosome and are 90% identical in DNA sequence. The
tps gene encodes the abundant spore coat protein S and its expression is activated about 5
hours after the initiation of development, whereas ops encodes a protein found inside
myxospores and is not expressed until much later in development, around the stage of
sporulation. A DNA segment between 131 and 311 bp upstream from the ops
transcriptional start site regulates not only the expression of ops, but also the expression
of tps located about 2 kb downstream. The activation is orientation independent and the
UAS is still functional when the DNA segment is moved farther upstream. The need for
an activator protein for tps expression is supported by the finding that a protein-DNA

complex was observed when an upstream DNA segment was incubated with extracts of

30

developing M xanthus cells. Also, expression of tps requires two upstream sequences
centered at -90 and -270 bp relative to its transcriptional start site.

The regulation of csgA gene expression provides another example of the likely
involvement of a transcriptional activator protein(s). Responding to environmental cues
such as nutrient level, peptidoglycan and B signal, csgA expression and the resulting
increase in ngA level during the developmental process is thought to entrain the order of
morphological events, i.e., ripping, aggregation, then sporulation (Li et al., 1992). Hence,
csgA expression must be tightly controlled. Indeed, the expression of csgA requires DNA
upstream of the core promoter. DNA extending to -—400 bp appears to be needed for
optimal csgA expression under starvation condition, whereas another 500 bp upstream
(up to —930 bp) is necessary for full expression under low nutrient level conditions.

C-signal-dependent Tn5 lac fusions Q4403 (Fisseha et al., 1996), Q4400
(Brandner and Kroos, 1998) and Q4499 (Fisseha et al., 1999) all require DNA upstream
of the region typically recognized by RNA polymerase for full developmental expression.
In the following chapter, I show that full expression of Q4514 also requires DNA
upstream of its core promoter sequence, suggesting the involvement of a transcriptional
activator protein.

I chose to study Q4514 regulation because it is expressed with similar timing
during development as the other Tn5 lac fusions just mentioned, but unlike the other
fusions, Q4514 expression does not depend on C-signaling. The promoters of the C-
signal-dependent genes are not similar in the —10 and —35 regions, and they do not

70 .
resemble the consensus sequence for c or 054 promoters. However, a C-rrch sequence

CATCCCT has been identified in the Q4403 and Q4400 promoters centered at —49.

31

Similar sequences have also been identified in the Q4499 promoter at position —55, and
in another C-signal-dependent promoter, csgA, at position —63. A degenerate consensus
S'-CAYYCCY-3' (Y means pyrimidine), known as the C box, has been deduced from
these sequences and it has been proposed that C-box sequences are cis-acting regulatory
elements important for C-signal—dependent gene expression (Fisseha et al., 1999). The
Q4514 promoter DNA does not contain a region similar to the C box sequence around —
50. Additionally, unlike C-signal-dependent promoters, the Q4514 promoter resembles
the promoters transcribed by (:70 RNAP and it can be transcribed by GA RNAP isolated
from vegetative M xanthus cells. These results suggest that Q4514 expression is
regulated by a different mechanism than C-signal-dependent genes. My investigation
revealed a new aspect of gene regulation during M xanthus development, thus
contributing to our overall understanding of developmental gene regulation in response to

cell-cell interactions.

32

Chapter II

Characterization of regulatory mechanism of a cell
interaction-dependent gene in M yxococcus xanthus

33

Abstract

Q4514 is a Tn5 lac insertion in the Myxococcus xanthus genome that fuses lacZ
expression to a developmentally regulated promoter. I cloned DNA upstream of the
Q4514 insertion site and localized the promoter. The promoter, which is induced during
development, resembles vegetative promoters in sequence. GA RNA polymerase, the
major form of RNA polymerase in growing M xanthus, initiated transcription from this
promoter in vitro. Two complete open reading frames were identified downstream of the
promoter and before the Q4514 insertion. The first gene product (ORFl) has a putative
helix-turn-helix DNA-binding motif and shows sequence similarity to several
transcriptional regulators. Our results show that the product of orfl exerts a negative
regulatory effect on expression of its own operon. However, we were unable to
demonstrate specific binding of ORF 1 to the Q4514 promoter region. The second gene
(0212) is most similar to subunit A of glutaconate CoA-transferase which is involved in
the glutamate fermentation pathway. Mild sporulation defects were observed in orfI and

orf2 disruption mutants, whereas the Tn5 lac Q4514 insertion did not cause a

developmental defect. Although ORF] negatively regulates Q4514 expression,
developmental induction of this operon does not result primarily from a loss of ORFl.
Rather, transcriptional activation appears to be involved and multiple upstream DNA
elements are necessary for full developmental expression. Q4514 is the first example of a
developmentally expressed M xanthus operon that is transcribed by the major vegetative
RNA polymerase, and its regulation appears to involve both negative autoregulation by

ORFl and positive regulation by one or more transcriptional activators.

34

Introduction

Myxococcus xanthus is a gram-negative soil bacterium that undergoes
multicellular development (Dworkin and Kaiser, 1993). When starved at high cell density
on a solid surface, cells move in a coordinated fashion into aggregation centers, where
they form mound-shaped fruiting bodies that each contain approximately 105 cells.
Within the fruiting bodies, some of the rod-shaped cells differentiate into dormant, ovoid
spores that are heat and desiccation resistant.

Cell-cell interactions play a critical role in this multicellular developmental
process. At least five extracellular signals, known as the A—, B-, C-, D-, and E-signals,
appear to be involved. Mutants defective in the production of any one of these signals are
arrested in development at a particular stage, but are rescued by codevelopment with wild
type cells or cells that are defective in the production of a different signal (Hagen et al.,
1978; LaRossa et al., 1983; Downard et al., 1984).

To study the role of cell-cell interactions in controlling gene expression during M
xanthus development, Tn5 lac, a transposon containing a promoterless E. coli lacZ gene,
has been used to identify developmentally regulated genes (Kroos and Kaiser, 1984).
Transposition of Tn5 lac into the M xanthus chromosome can generate a transcriptional
fiision between lacZ and an M xanthus promoter. Among 23 74 Tn5 lac insertions, 29 of
them were activated during M xanthus development (Kroos et al., 1986). The
dependence of developmental gene expression on cell-cell interactions was examined by
monitoring B-galactosidase expression of the lacZ fusions in cell interaction mutants

(Kroos et al., 1986). A- and B-signaling are required for normal developmental gene

35

expression at the onset of development (Gill and Cull, 1986; Kuspa et al., 1986; Kroos
and Kaiser, 1987), D- and E-signaling are required at 3 to 5 hr into development (Cheng
and Kaiser, 1989b; Downard et al., 1993), and C-signaling is required at about 6 hr into
development (Kroos and Kaiser, 1987; Li and Shimkets, 1993).

Among the five extracellular signaling interactions, only the A- and C-signaling
interactions are well-characterized. A-signal is a mixture of peptides and amino acids
apparently generated by extracellular proteases released in response to nutrient limitation
and used to determine whether cells are at a sufficiently high density to initiate
multicellular development (Plamann et al., 1992; Kuspa et al., 1992; Kuspa et al., 1992).
All of the C-signaling mutations map to a single gene called csgA, and the mutants are
defective in both fi‘uiting body formation and sporulation (Hagen et al., 197 8; Shimkets
and Kaiser, 1982; Shimkets and Asher, 1988). ngA, a cell surface and/or extracellular
matrix-associated protein (Shimkets and Rafiee, 1990), appears to mediate C-signaling,
which both controls cell movements (Sogaard-Anderson and Kaiser, 1996; Jelsbak and
Sogaard-Anderson, 1999) and is influenced by the alignment of cells that results from
movement of cells into aggregates (Kim and Kaiser, 1990a; Sager and Kaiser, 1994).
Close packing of cells within fiuiting bodies is thought to allow efficient C-signaling,
which triggers sporulation (Sager and Kaiser, 1994). Different levels of C-signaling are
also required for expression of different developmental genes (Kim and Kaiser, 1991).
Hence, C-signaling seems to act as a timer governing the developmental process and
appears to couple morphogenesis of the fruiting body with expression of genes at the

proper times.

36

We are interested in the regulation of developmental gene expression in response
to extracellular signals. To begin to understand the mechanisms involved, the regulatory
regions of several C-signal-dependent transcriptional units, identified by Tn5 lac
insertions Q4403, Q4400 and Q4499, have been characterized (Fisseha et al., 1996;
Brandner and Kroos, 1998; Fisseha et al., 1999). The promoters are not similar to each
other in the -10 and —35 regions and they do not resemble promoters that are transcribed
by O70 or 0'54 RNAP. However, both Q4403 and Q4400 have the sequence 5'-
CATCCCT-3' centered at —49 bp. Similar sequences are also found in the Q4499
promoter region and other C-signal-dependent promoters at positions near --50 bp.
Changing 5’-CATCCCT-3' in the Q4400 promoter to 5'-ACGAAAG-3’ abolished
activity of this promoter, demonstrating the importance of this sequence. Hence, a
sequence with the consensus 5 '-CAYYCCY-3' (known as the C box) has been proposed
to be important for C-signal-dependent gene expression.

Here, I report studies on the regulation of Q45 14, a Tn5 lac insertion that is
expressed with similar timing during development as Q4400, Q4403 and Q4499, but
Q4514 expression does not depend on C-signaling. I constructed a series of 5' and 3’
deletions to localize the sequence required for promoter activity, determined the
nucleotide sequence of the promoter region, and mapped the transcriptional start site. It
appears that the Q4514 promoter is different from other M xanthus development-specific
promoters that have been characterized in that it resembles (:70 promoters and can be
transcribed by GA RNAP isolated from vegetative M xanthus cells. Also, no sequence
matching the C box consensus is present near —50 bp in this C-signal—independent

promoter. Regulation of the Q4514 operon is complex, involving negative autoregulation

37

by the product of the first gene of the operon and positive regulation mediated by
upstream DNA elements. These studies provide the first example of a development-
specific M xanthus gene that is transcribed by the major vegetative RNAP and establish
a foundation for biochemical approaches toward identifying proteins that regulate Q4514

expression.

38

Materials and Methods

Bacterial strains and plasmids. Strains and plasmids used in this work are listed

in Table 2.1.

Growth and development. Escherichia coli cells were grown at 37°C in LB
medium (Sambrook et al., 1989) containing 50 ug ampicillin, 25 ug kanamycin or 10 ug
of tetracycline per ml, as necessary. M xanthus was grown at 32°C in CTT medium
(Hodgkin and Kaiser, 1977) in liquid culture or on agar (1.5%) plates with 40 ug of

kanamycin or 12.5 pg of oxytetracycline per m1 when required (Kroos et al., 1986).
Fruiting body development was performed on TPM (10 mM Tris-HCl, pH 8.0, 1 mM
KH2P04, 8 mM MgSO4, final pH 7.6) agar (1.5%) plates as described previously (Kroos
et al., 1986).

Molecular cloning. Recombinant DNA work was performed using standard
techniques (Sambrook et al., 1989). Plasmid DNA was prepared from E. coli DHSa or
JMS3.

To clone the DNA upstream of Tn5 lac Q4514, chromosomal DNA was prepared
(Laue and Gill, 1994) from M xanthus DK4514 and digested with XhoI, the fragments
were ligated to XhoI-digested pGEM-7Zf, and the mixture was transformed into E. coli
DHSa, selecting for both ampicillin resistance (Ap') of the vector and kanamycin
resistance (Km’) of the desired insert. One transforrnant with a plasmid bearing an insert
of the expected size was characterized fiirther. Restriction fragments of M xanthus DNA
from this plasmid, pTH1-3, were gel-purified and ligated into vectors as indicated in

Table 2.1. In these and the subsequent subcloning steps described in Table 2.1, vectors

39

Table 2.1. Bacterial Strains and Plasmids

 

Strain or plasmid

Characteristics

Source or Reference

 

E. coli

DH50t

JM83

BL21(DE3)

M xanthus

DK1622

DK4514

MMFl727

MTH1-3
MTH4-1
MTH4-2
MTH4-3
MTH5-3
MTH5-4
MTHS-S
MTH10-3

MTHll-l

supE44 AlacU169 (1)80 AlacZ AMIS 11st17

recAl endAl gyrA thi-l relAl

ara Alac-pro sirA (hi ¢dlacZ AM15

F‘ ompT gal [dcm] [Ion] hstB (r3°m3';an
E. coli B strain) with DE3, a X prophage carrying

the T7 RNA polymerase gene

wild type

Tn5 lac (Kmr) 04514

attB :

atth

atth

attB:

atth

atth

attB :

atlB:

attB:

atiB:

:pREG1727

:pTI-Il-6
:pTH4-1
:pTH4-2
:pTH4-3
:pTH5-3
:pTH5-4
:pTH5-5
:pTH10-3

:pTH10-3 orflszI-Il 1-1

40

Hanahan,
1 983

Messing, 1979

Novagen

Kaiser, 197 9

Kroos et al., 1986

Fisseha et al.,
1996

This study
This study
This study
This study
This study
This study
This study
This study

This study

MTHl 1-2 0112 : pTHl 1-1 This study

MGVOI orf1::pBRE47Sp4514 Greg Velicer
MGV02 attB::pTH10-3 orfI ::pBRE47Sp4514 Greg Velicer
MTH8-1 Tn5 lac (Tor) Q4514 This study
MTH9-1 MTH8-l ::pTH9-1, Kmr Tcr This study
MTH9-2 MTH8-l ::pTH9-2, Kmr Tcr This study
MTH9-4 MTH8-l ::pTH9-4, Kmr Tcr This study
DK5208 csgA::Tn5-132 (Tc') Q205 Shimkets and
Asher, 1988
MTH7-1 csgAzzTn5-132 (Tc') Q205 atthszH1-6 This study
MTH7-2 csgAzzTn5-132 (Tc') Q205 attB::pTH4-3 This study
MTH7-3 csgA12Tn5-132 (Tc’) Q205 attB::pTH4-1 This study
JPB03 csgAzzTn5-132 (Tc’) Q205 attB::pREG1727 Brandner and
Kroos, 1998

DK5225 csgA::Tn5-132 (Tc’) Q205 Tn5 lac (Kmr) Q4514 Kroos and

Kaiser, 1987
MGV03 DK1622::p1 175 SmSl45 14 Greg Velicer
Plasmid
pUC19 Apr laca Yanisch-Perron et al.,

1985

pTH4-4 Apr(pUC19)“; 1.6-kb SmaI-BamI-II fragment from pTH1-3 This study
pTH4-6 Apr (pUCl9); 0.7-kb SacI-BamHI fragment from pTH4-4 This study
pTH4-7 Apr (pUCl9); 1.3-kb SphI-BamHI fragment from pTH4-4 This study
pTH7-1 Apr (pUCl9); 2.2-kb SalI-BamHI fragment from pTH1-3 This study

41

pTH7-2 Apr (pUCl9); 4.3-kb )flroI-BamHI fi'agment from pTHl-3 This study

pTHlO-l Apr (pUCl9); 120-bp SmaI-Eco47III fragment from pTH4—4 This study

pGEM-7Zf Apr laca Promega

pTH1-3 Apr Kmr (pGEM-7Zf); 13.3-kb XhoI fragment This study
from DK4514

pTHS-l Apr (pGEM-7Zf); 0.7-kb EcoRI-BamI-II fragment fi'om This study
pTH4-6

pTH5-2 Apr (pGEM-7Zf ); 1.3-kb NruI-BamI-II fragment from This study
pTH4-4

pGEME47Sp4514 Apr (pGEM-7Zf); 242-bp Eco47III-SphI fragment This study

from pTH4-4

pTH9-3 Apr (pGEM-7Zf); 4.3-kb HindIII-BamI-II fragment fiom This study
pTH7-2

pBR322 Apr Tcr Bolivar et al., 1977

pBRE47Sp4514 Tcr (pBR322); 250-bp NsiI-AatII fragment from This study

pGEME47Sp4S 14
pTHl 1-1 Tcr (pBR322); 323-bp fragment of internal or]? This study
sequence generated by PCR of pTH4-4
pET2 1 a Apr Novagen
pTH191 Apr (pET21a); complete orfl sequence generated by This study
PCR from pTH4-4 using primers LK190 and LK191
pTHl92 Apr (pET21a ); complete orfl sequence followed by This study
a stop codon generated by PCR from pTH4-4 using
primers LK190 and LK192
pET16b Apr Novagen
pTH193 Apr (pET16b); complete orf1 sequence generated by This study

PCR from pTH4-4 using primers LK190 and LK193

42

pREG1727 Apr Kmr Pl-inc attP lacZ Fisseha et al.,1996

pTH1-6 Apr Kmr (pREG1727); 4.3-kb XhoI-BamHI fi'agment This study
from pTHl-3

pTH4-1 Apr Kmr (pREG1727); 2.2-kb SalI-BamI-II fragment This study
from pTH1-3

pTH4-2 Apr Kmr (pREG1727); 2.1-kb XhoI-SalI fragment This study
from pTH1-3

pTH4-3 Apr Kmr (pREG1727); 1.6-kb SmaI-BamHI fragment This study
from pTH1-3

pTH5-3 Apr Kmr (pREG1727); 0.7-kb XhoI-BamHI fragment This study
from pTHS-l

pTH5-4 Apr Kmr (pREG1727); 1.3-kb XhoI-BamI-II fi'agment This study
from pTH5-2

pTH5-5 Apr Kmr (pREG1727); 1.3-kb HindIII-BamHI fragment This study
from pTH4-7

pTH10-3 Apr Kmr (pREG1727); 120-bp HindIII-BamI-II fragment This study
from pTHlO-l

pREG429 Apr Kml' Pl-inc Gill et al.,1988

pTH9-1 Apr Kmr (pREG429); 1.6-kb EcoRI-BamHI fragment This study
from pTH4-4

pTH9-2 Apr Kmr (pREG429); 2.2-kb Ach-BamHI fragment This study
from pTH7-1

pTH9-4 Apr Kmr (pREG429); 4.3-kb EcoRI-BamHI fragment This study
from pTH9-3

pREGl 175 Apr Kmr Pl-inc Gill and

Bornemann, 1988

pl 17SSmSl4514 Apr Kmr (pREG1175); 2.7-kb XhoI-SmaI fragment Greg Velicer
from pTH7-2

 

a The vector is indicated in parentheses.

43

were digested with the same restriction enzymes used to produce the fi'agments, except as
indicated below.

To construct pTH5-2, the 0.9-kb NruI-BamI-II fragment from pTH4-4 was gel-
purified and ligated into pGEM-7Zf that had been digested with Smal and BamHI.

Plasmid pTH9-2 was constructed as follows. The 2.2-kb Ach-BamHI fragment
from pTH7-1 was gel-purified and ligated into pREG429, which had been digested with
C laI and BamHI.

To subclone the 120-bp SmaI-Eco47III fragment from the region upstream of Tn5
lac Q4514 into pREG1727, the fragment was gel-purified after digestion of pTH4-4 and
ligated into SmaI-digested pUC19. A plasmid with the fragment in the correct orientation
for further cloning into pREG1727 (i.e., the SmaI end of the fragment close to the Hindlll
site in pUC19) was identified and named pTHlO-l. A HindIII-BamI-II fragment of
pTHlO-l that contains the 120-bp SmaI-Eco47III fragment of Q45 14 was obtained and
cloned into HindlII-BamHI-linerized pREG1727, resulting in pTH10-3, in which the 120-
bp SmaI-Eco47III fragment was positioned before the promoterless lacZ.

Plasmid pGEME47Sp4514 was constructed by digestion of pTH4-4 with
Eco47III and SphI, gel-purification of the resulting 242-bp fragment and ligation into
pGEM-7Zf, which had been linerized with SmaI and SphI. This plasmid was digested
with NsiI and AatII to obtain a 250-bp fragment, which was then ligated into AatII-Pstl-
digested pBR322 to construct pBRE47Sp4514.

Plasmid pTHl 1-1 was constructed by ligating part of the 0272 sequence into
pBR3 22. The partial orf2 sequence was generated by PCR using pTH4-4 as a template.

The upstream primer was 5'-AAACTGCAGTCCGGATGGCGCGTCG-3', which is

44

shown with Pstl site underlined. This primer anneals to the sequence between positions
814 and 829 (Figure 2.4), and the downstream primer was 5'-
AAGAATTCGGGATGAAGGGCAGCC-3', which has an underlined EcoRI site and
anneals to the sequence between positions 1137 and 1122 (Figure 2.4). The amplified
fragment was digested with Pstl and EcoRI, gel-purified, and ligated to Pstl-EcoRI-
digested pBR3 22 to construct pTHl 1-1. The insert was sequenced to ensure that no error
occurred during the PCR.

Plasmids pTH191, pTHl92 and pTH193 were constructed as follows. The insert
for each plasmid was generated by PCR with the same upstream primer, 5'-
AGAAGGGACATATGACGAACACCGGAGGA-3', which has an NdeI site
(underlined) and the first 18 bases of orf1 sequence. The downstream primer (LK191) for
generating the insert of pTHl91 was 5'-CTCCTCGAGTGCGTCCTCCGAATCCGT-3',
which has a XhoI site (underlined) and the last 18 bases of orfI sequence without a stop
codon. The downstream primer (LK192) for generating the insert of pTHl92 was 5'-
CTCCTCGAGTCATGCGTCCTCCGAATC-3', which has the stop codon TGA adjacent
to the X7201 site. The downstream primer (LK193) for generating the insert of pTH193
was like LK192, except it has a BamHI site instead of the XhoI site. The amplified
fragments were digested with appropriate restriction enzymes, gel-purified and ligated
into NdeI-XhoI-digested pET21a to construct pTH191 and pTHl92, or into NdeI-BamHI-
digested pETl6b to construct pTH193. Plasmids pET21a and pETl6b are designed for
high level protein expression in E. coli under the control of T7 RNA polymerase. The
01f] DNA was upstream of an in-frame sequence encoding a 6xHis tag followed by a

stop codon in pTH191, while in pTHl92, the orfl DNA was followed immediately by a

45

stop codon. In pTH193, the orf1 DNA was downstream of an in-fiame sequence
encoding a 6xHis tag. The inserts and junctions were sequenced to ensure that no error
occurred during the PCR

To construct p1175SmSl4514, the 2.7-kb )flroI-Smal fragment from pTH4-4 was
gel-purified and ligated into pREG1175 that had been digested with Sall and SmaI.

DNA sequencing. Plasmid pTH4-4 was used as the template in sequencing
reactions performed by the method of Sanger et al. (Sanger et al., 1977) with a Sequenase
kit (United States Biochemical). Ambiguities arising from premature termination were
resolved using the protocol of Fawcett and Bartlett (F awcett and Bartlett, 1990). Briefly,
lpl of a reaction mixture containing terminal deoxynucleotide transferase (1 pM each
deoxynucleoside triphosphate [pH 7 .0], 2 U of terminal deoxynucleotide transferase per
u], 1x Sequenase reaction buffer) was added to each of the termination reactions (16 Ill)
and incubated at 37°C for 30 min. The reaction was terminated by adding 4 pl of stop
buffer (United States Biochemical). 7-Deaza-dGTP reaction mixtures were used to
resolve regions of compression. DNA and protein sequence analyses were done with the
University of Wisconsin Genetics Computer Group software package. The Michigan
State University Macromolecular Structure Facility synthesized oligonucleotide primers
as needed to sequence both strands of the 1.9-kb Q4514 upstream DNA.

Construction of M xanthus strains. Strains containing pREG1727 derivatives
integrated at Mx8 attB were constructed by P1 specialized transduction from the rec+ E.
coli strain JM83 (Gill et al., 1988; Fisseha et al., 1996), or by electroporation, of the wild-
type M xanthus strain DK1622 or the csgA mutant strain DK5208. Except where noted

otherwise, at least three derivatives, each containing a single copy of integrated plasmid,

46

were identified by Southern blot analysis (Sambrook et al., 1989; Fisseha et al., 1996),
and B-galactosidase production was measured under developmental conditions as
described previously (Kroos et al., 1986).

Strain MTHS-l was constructed by transducing bacteriophage P1::Tn5 lac (Tc’)
into DK4514 with selection for oxytetracycline resistance. Screening for kanamycin-
sensitive transductants identified MTH8-1 in which the Kmr gene was replaced by the Tcr
gene, as verified by Southern blot analysis (Brandner and Kroos, 1998). MTH9-l,
MTH9-2 and MTH9-4 are Kmr Tcr strains resulting from P1 specialized transduction of
pTH9-l, pTH9-2 and pTH9—4 from E. coli JM83 into M xanthus MTH8-1, respectively.

MGVOI and MTHl 1-1 were constructed by electroporating pBR322 derivatives
pBRE47Sp4514 and pTHl 1-1 into DK1622 and MTH10-3, respectively. Three
transforrnants containing a single copy of either plasmid integrated by homologous
recombination at orfl or my? were identified by Southern blot analysis and PCR (data not
shown). Likewise, MGV03 was constructed by electroporating p1175SmSl4514 into
DK1622.

Polymerase chain reaction. Diagnostic PCR was used to identify M xanthus
integrants as follows. Cells were grown in CTT medium with appropriate antibiotics,
collected by centrifugation, and resuspended in 500 pl of TE with 100 jig/ml RNAse A.
The cell suspension was incubated first at room temperature for 15 min and then at 85°C
for 15 min, followed by extraction with one volume of phenol, then two extractions with
one volume of chloroform. The aqueous supernatant after microcentrifugation was used

for the PCR reaction.

47

RNA analysis. RNA was prepared as described previously (Fisseha et al., 1996)
from M xanthus DK1622. An S1 nuclease protection assay was preformed with a probe
from pTH7-2 digested with NruI. The 2.6-kb fragment was gel-purified, phosphatase-
treated, and 5' end-labeled with [7-32P] ATP and T4 polynucleotide kinase. RNA (100 ug)
was precipitated with the labeled probe. The pellet was resuspended in 30 ul of
hybridization buffer (80% formamide, 20 M pipes [pH 6.4], 0.4 LIM NaCl), and
incubated for 10 min at 85°C followed by 3 hr at 52°C. 81 nuclease buffer (0.03 M
NaOAc [pH4.6], 0.05 M NaCl, 1 mM ZnSO4, 5% glycerol) and 500 u of S1 nuclease
(Boeringer Mannheim) were then added, giving a final volume of 300 pl. After 30 min at
37 °C, the reactions were extracted with 300 pl phenol-chloroform, precipitated with
ethanol, and resuspended in formamide loading buffer (80% formamide, 10 mM EDTA,
0.01% xylene cyanol, 0.01% bromophenol blue). The protected products were resolved
on a 5% polyacrylamide-8M urea gel and visualized by autoradiography.

Primer extension analysis was performed as described previously (Sambrook et
al., 1989), using the oligonucleotide 5'-GATCTCCTGCATCGACGTGCCCTC-3', which
corresponds to a sequence located about 140 bp downstream of the mRNA 5' end
mapped by S1 nuclease protection. The primer was end-labeled with T4 polynucleotide
kinase and [7-32P] ATP and purified with a QIAquick Nucleotide Removal Kit (Qiagen).
The reaction products were analyzed on a 5% polyacrylamide —8 M urea sequencing gel
next to dideoxy sequencing reactions that utilized the same primer.

In vitro transcription. Reactions were performed with partially purified M
xanthus oA RN AP or reconstituted O'A RNAP holoenzyme as described previously (Biran

and Kroos, 1997).

48

Western blot analysis. E. coli BL21 (ADE3) containing pTHl92 was grown in
LB medium containing ampicillin and harvested by centrifugation 3 h after 0.4 mM [PTG
induction. The cell pellet was resuspended in 1x sample buffer (0.125 M Tris-HCl pH
6.8, 5% B-mercaptoethanol, 2% SDS, 10% glycerol, 0.1% bromophenol blue) and boiled
for 5 min. DK1622 and orfl mutant (MGVOI) cells were collected at the indicated times, I
resuspended in sample buffer, and boiled for 5 min. Equal amounts of M xanthus protein,
as determined by measuring total protein in sonic lysates of the same samples by the
Bradford method (Bradford, 1976), were subjected to Western blot analysis. Proteins
were separated by 10% or 12% SDS-PAGE and electrotransferred to a nitrocellulose or
poly(vinylidene difluoride) membrane. The membrane was incubated in TBST (20 mM
Tris-HCl pH 7.5, 0.5 M NaCl, 0.1% Tween 20) containing 2% nonfat dry milk for 4 h at
room temperature with shaking in order to block nonspecific interaction between the
primary antibodies and the membrane. The membrane was then probed for 4 h with
shaking at room temperature with either a polyclonal antiserum to B. subtilis 0’43 (a gift
from B. —Y. Chang and R. Dori) or affinity-purified anti-ORF1-6xHis antibody diluted
1:500 in TB ST/ 2% nonfat dry milk. Irnmunodetection using goat anti-rabbit-IgG
horseradish peroxidase (Bio-Rad) and chemiluminescence (ECL; Amersham Pharmacia
Biotech) was performed according to the manufacturer's instructions.

Production of ORFl-6xHis. pTH191 was transformed into E. coli strain
BL21(XDE3) (N ovagen), which contains the gene for T7 RNA polymerase under the
control of an [PTG-inducible lacUV5 promoter. Cells containing pTH191 were
inoculated into LB medium containing ampicillin and grown at 37 °C until the ODsoo

reached about 0.6, then the culture was induced with 0.4 mM IPTG. Cells were collected

49

3 hours after the induction by centrifiigation (5,000xg, 5 min) and the cell pellet was
stored at -70°C. Some experiments suggested that the ORF1-6xHis might be located in
cytoplasmic inclusion bodies (data not shown). To minimize the proportion of ORFl-
6xHis incorporated into inclusion bodies, cells were grown at 30°C and induced with
IPTG as described above.

A whole-cell extract of the IPTG-induced cells was prepared as follows. The cell
pellet was resuspended in 10 ml of 5 mM imidazole, 0.5 M NaCl, 20 mM Tris-Cl pH 7.9,
1 mM PMSF. Cells were lysed by sonication (Sonicator W-225; microtip maximum
power setting at 6; Heat System-Ultrasonics, Inc.) while the mixture was kept cold on ice.
The lysate was centrifiiged at 39,000xg for 20 min at 4°C.

The ORF1-6xI-Iis protein in the supernatant was isolated under nondenaturing
conditions using a Ni-column as follows. The well-packed Ni-column was washed with 5
volumes of 1x column binding buffer (5 mM imidazole, 0.5 M NaCl, 20 mM Tris-Cl pH
7.9) and allowed to drain until the buffer reached the top of the column bed. After the
whole-cell extract was loaded onto the column, it was washed with 10 volumes of 1x
column binding buffer, followed by 6 volumes of 1x wash bufi‘er (60 mM imidazole, 0.5
M NaCl, 20 mM Tris-Cl pH 7.9). Proteins were then eluted with 6 volumes of 1 M
imidazole, 0.5 M NaCl, 20 mM Tris-Cl pH 7.9. The eluate was dialyzed against 2 L of
gel-shift binding buffer (12% glycerol, 20 mM HEPES [pH 7 .9], 50 mM KCl, 1 mM
EDTA, 1 mM DTT, 5 mM MgC12) for 16 hr at 4°C. The protein solution (around 30 ml)
was then concentrated 20-fold by centrifugation in a Millipore 15 filter device for 3 h at

4°C using the manufacturer's specifications and stored at 4°C.

50

Mobility shift experiments. The SmaI-Eco47III fragment containing DNA from
—55 bp to +65 bp relative to the transcriptional start site was used as probe in the mobility
shift experiments. This fragment from pTH4-4 digested with SmaI and Eco47III was gel-
purified, phosphatase-treated, and 5' end-labeled with [7-32P] ATP and T4 polynucleotide
kinase. The labeled DNA was purified with a QIAquick Nucleotide Removal Kit
(Qiagen). Various amounts of affinity-purified ORF1-6xHis protein or crude extracts
made from cells overexpressing ORF1-6xHis, ORF] or 6xHis-ORF l were incubated with
4.5ng (57 frnol) probe and 1 pg poly(dI'dC) at 30°C for 15 min to allow DNA binding.
The 20 ul binding reactions contained 12% glycerol, 20 mM HEPES (pH 7.9), 50 mM
KCl, 1 mM EDT A, lmM DTT, 5 mM MgC12. Samples were then subjected to
electrophoresis on a 5% or 8% polyacrylamide gel in low-ionic strength buffer at 4°C
(Chodosh, 1988). After electrophoresis, the gels were dried and exposed to film.

Preparation and purification of antibodies. Purified ORF1-6xHis protein (500
ug) was mixed with Freund's complete adjuvant (BRL) and injected subcutaneously into
a rabbit. Four weeks later, a booster injection (3 00 ug of ORF1-6xI-Iis mixed with
Freund's incomplete adjuvant) was performed. The rabbit was bled 1 week after the boost
and the serum was prepared as described previously (Harlow and Lane, 1988).

Affinity purification of the antibodies was performed as follows. Purified ORF1-
6xHis protein (100 pg) was electrophoresed on a 12% polyacrylamide-SDS gel and the
protein was electrotransferred to a poly (vinylidene difluoride) (PVDF) membrane
(Millipore). The region of the membrane with bound ORF1-6xHis was excised and
incubated with TBST (20 mM Tris-HCl pH 7.5, 0.5 M NaCl, 0.05% Tween 20)

containing 5% nonfat dry milk for 2 h to block non-specific binding of the antibodies.

51

Anti-ORF1-6xHis antiserum (2 ml) was incubated with the membrane overnight at 4°C
with agitation. The membrane was then washed twice (5 min each) with TB ST and twice
with TBS (20 mM Tris-HCl pH 7 .5, 0.5 mM NaCl). To elute antibodies bound to the
membrane, 100 mM glycine pH 2.5 (1 ml) was added and incubated for 10 second with
shaking. The solution was collected and immediately neutralized to pH 7.0 by adding 1
M Tris-Cl pH 8.0 (approximately 100 [4.1). The elution steps were repeated twice and the
neutralized solutions were combined and stored at 4°C.

Sporulation efficiency. Cells were plated for development on TPM agar (1.5%)
and harvested after 3 days at 32°C, then the samples were subjected to heat and
sonication treatment to kill rod-shaped cells, and dilutions were plated to quantitate the
number of spores able to germinate and form colonies, as described previously (Kroos

and Kaiser, 1987).

52

Results

Localization of the developmental promoter controlling Q4514 expression.
To clone the putative promoter located upstream of the developmentally regulated Tn5
lac insertion Q4514, I took advantage of a X7101 restriction site approximately 4.3 kb
upstream of the Q4514 insertion in M xanthus DK4514 (Kroos et al., 1986) and a XhoI
site in Tn5 lac approximately 9 kb from the left end (Figure 2.1). Chromosomal DNA
from DK4514 was digested with XhoI, cloned into pGEM-7Zf, and transformed into E.
coli cells. Since the 13.3 kb X7101 fragment described above includes the aphII gene of
Tn5 lac, which encodes aminoglycoside phosphotransferase and confers Km’, E. coli
cells that contain plasmids with the desired XhoI fiagment of M xanthus chromosomal
DNA will survive under Km selection. Several restriction sites upstream of the Q4514
insertion in DK4514 have been mapped by Kroos et al. (Kroos et al., 1986). Restriction
mapping of the resulting plasmid, pTH1-3, showed the patterns expected on the basis of
restriction sites in DNA upstream of Q45 14 and on the basis of known restriction maps
of pGEM-7Zf and Tn5 lac. Figure 2.1 shows a restriction map of DNA upstream of
Q4514 generated on the basis of these results.

To test Q4514 upstream DNA for promoter activity, the XhoI-BamHI restriction
fragment from pTH1-3, which contains 4.3 kb of M xanthus DNA and approximately 60
bp of the left end of Tn5 lac (Figure 2.1), was subcloned into XhoI-BamI-[I-digested
pREG1727 to construct pTH1-6. Since the BamI-II site of pREG1727 is located
immediately upstream of the same lacZ-containing fragment found in Tn5 lac (F isseha et

al., 1996), pTH1-6 contains Q4514 upstream DNA fused to a promoterless lacZ gene in

53

Figure 2.1. Physical map of the Q4514 insertion region and summary of deletions
tested for promoter activity. The upper schematic depicts the restriction sites in Tn5 lac
and the upstream M xanthus chromosome that were used in this study. Distances of
restriction sites from the Tn5 lac Q4514 insertion are given in kilobases. B, BamHI; E,
E004 7111; N, NruI; Sc, SacI; S1, Sall; Sm, SmaI; Sp, SphI; X, A7101. The lower diagram
depicts the segments of Q45 14 upstream DNA fused to the promoterless lacZ gene in
pREGl727 or in pREG1175 in the case of strain MGV03, to test for promoter activity.
The plasmid or strain designation is indicated on the left (Table 1). Derivatives of wild-
type M xanthus DK1622 containing a single copy of each pREGl727 derivative
integrated at Mx8 attB or a single copy of the pREG1175 derivative integrated by
homologous recombination were measured for B-galactosidase. The maximal l3-
galactosidase specific activities during a 72-hr developmental time course are given as
percentages of the maximum activity observed for Tn5 lac Q4514-containing strain
DK4514. In each case, the activity of at least three independent transductants was
measured at least once as described previously (Kroos et al., 1986) and the average is

given.

54

? lacZ aphll )f

—>—>
X 81 Sm E N Sp Sc\/
|—\\\ 1 1 . I" 1 .

 

 

 

 

 

 

 

 

! ' \
4.3 2.2 1.621.50 1.29 1.26 0.7 o
fi-galactosidase

Activity
pTH1-6 I ' ‘ ' 66%
pTH4-1 I I 37%
pTH4-3 I J. 16%
pTH10-3 LJ 14%
pTH5-4 I I 1%
pTH5-5 I J 1%
pTH5-3 I . 1%
MGV03 l J 1%

 

55

the same manner as Q4514-containing M xanthus DK4514. pTH1-6 was transduced
from E. coli JM83 into the wild-type M xanthus strain DK1622 using bacteriophage P1
specialized transduction (Fisseha et al., 1996). Due to the presence of the attP segment
from myxophage Mx8, pTH1-6 integrated efficiently into the M xanthus chromosome at
attB (Stellwag et al., 1985; Stephens and Kaiser, 1987). Transductants containing a single
copy of pTH1-6 integrated at Mx8 attB were identified by Southern blot hybridization
(data not shown). Several of these transductants were assayed for B-galactosidase activity
during development and showed, on average, 66% of the maximal level observed in
DK4514 (Figures 2.1 and 2.2). The results indicate that the 4.3 kb Q4514 upstream DNA
segment contains a promoter(s) that is able to direct development-specific expression.

To further localize the developmental promoter(s) within this region, additional
deletions of the Q4514 upstream DNA were generated and tested for promoter activity as
described above. The results showed that the 2.2-kb and 1.62-kb Q4514 upstream DNA
segments directed lacZ expression with a similar timing as the Q4514-containing strain,
but reached about 37% and 16% of the maximal level, respectively (Figures 2.1 and 2.2),
however, transductants containing the 1.29-kb, 1.26-kb, or 0.7-kb Q4514 upstream DNA
segments produced no developmental B-galactosidase above the background (Figure 2.1).
These results indicate that a development-specific promoter driving the expression of Tn5
lac Q4514 lies between the SmaI and NruI sites, and that DNA farther upstream also
contributes significantly to promoter activity. To test for an additional promoter upstream
of the SmaI site, the 2.7-kb XhoI-SmaI fragment was inserted into pREG1175 (Gill and
Bornemann, 1988), resulting in p117SSmSl4514, in which the XhoI-Smal fragment is

fiised to the same promoterless lacZ-containing fragment found in Tn5 lac. Since

56

Figure 2.2. Developmental expression of lacZ under the control of the Q4514 promoter.

Developmental B-galactosidase specific activity was measured as described previously

(Kroos et al., 1986) for Tn5 lac-containing strain DK4514 (O) and for at least three

independent isolated transductants of DK1622 containing a single copy of the 4.3-kb (I),

2.2-kb (A), or 1.62-kb (1]) Q4514 upstream DNA fused to promoterless lacZ within
pREGl727 and integrated at Mx8 attB. The B-galactosidase specific activity of DK1622
containing the pREGl727 vector alone with no insert integrated at Mx8 attB (A) is also

shown. The average B-galactosidase specific activity from three determinations for
DK4514 and from at least one determination for each of three independent transductants
is plotted. B-galactosidase specific activity is expressed in nanomoles of o-nitrophenyl

phosphate per minute per milligram of protein.

57

 

 

 

 

400 -

p F r p — E

0 0 0 0 0 0 0 0
5 0 5 0 5 0 5

3 3 2 2 1 4|

>=>=om 050on 0mmo_m9.om_mm-u

20 so 40 so 60 7o 80
Time (Hr)

1O

58

pREG1175 does not contain the attP segment of Mx8, when p117SSmSl4514 was
transduced into wild-type DK1622, homologous recombination between the plasmid M
xanthus DNA segment and the M xanthus chromosome resulted in MGV03, in which the
lacZ reporter is positioned immediately downstream of the SmaI site in the chromosome.
No developmental promoter activity was observed in three such independent
transductants (Figure 2.1). We conclude that there is no additional promoter upstream of
the SmaI site that can function in the absence of downstream DNA to drive Tn5 lac
Q4514 expression.

Localization of an mRNA 5’ end upstream of Q4514. To test whether an
mRNA 5’ end maps upstream of the Q4514 insertion, an S] nuclease protection assay
was performed with a probe labeled at the Nrul site located 1.29-kb upstream of Q45 14
(Figure 2.1) and including about 2.6-kb of M xanthus DNA farther upstream. The probe
was hybridized to RNA from M xanthus DK1622 and subjected to S1 nuclease digestion.
The probe protected a development-specific RNA species as shown in Figure 2.3. RNA
from developing cells harvested at 12 or 18 h after starvation protected a fragment of
about 270 bases in length. No protection was observed with RNA from growing cells and
very weak protection was observed with RNA from 24-h developing cells. These results
indicate that a development-specific mRNA is transcribed from the region upstream of
the Q4514 Tn5 lac insertion and the 5’ end of this mRNA species is located
approximately 1.56 kb upstream of the insertion site. Together with the deletion analysis
(Figures 2.1 and 2.2), the results suggest that a promoter lies 1.56 kb upstream of Q45 14

and the mRNA 5' end reflects the transcriptional start site.

59

Figure 2.3. Localization of an mRNA 5' end within the Q4514 upstream region. Low-
resolution S1 nuclease mapping of the Q4514-associated transcript was

performed by using RNA prepared from DK1622 cells growing vegetatively (lane 1) or
cells that had undergone 12, 18, and 24 hr of development in lanes 2 to 4, respectively.
The numbers on the left indicate the length (in bases) of end-labeled MspI-digested

pBR3 22 marker fragments. The arrowhead denotes the protected fragment.

6O

 

61

DNA sequence of the Q4514 upstream region. The nucleotide sequence of both
strands of the 1.9-kb DNA fragment immediately upstream of the Q4514 insertion site
was determined (Figure 2.4). Two complete open reading fiames (ORFs) and one partial
ORF interrupted by Q4514 were inferred from the sequence downstream of the Q4514
transcriptional start site. The two ORFs exhibit codon preference typical of M xanthus
genes, including a strong bias towards usage of guanine or cytosine at the third codon
position (Shimkets, 1993). The first ORF (ORFl), beginning with GTG at position 434
and ending with a TGA stop codon at position 1118, is predicted to encode a 228 amino
. acid polypeptide. The second ORF (ORF2), beginning at position 1117 and ending at
position 1909, is predicted to encode a 264 amino acid polypeptide. A putative third
ORF, beginning at position 1908, is interrupted by Q4514 insertion after the first two
amino acids. Each open reading frame is preceded by a sequence that might serve as a
ribosomal binding site (AGAAGGGAG 5 bp upstream of ORFl, GGAGG 4 bp upstream
of ORF2 and GGAGGG 4 bp upstream of ORF3), since it is complementary to the
sequence near the 3' end of M xanthus 16S rRNA (Oyaizu and Woese, 1985). The
positions of the start points of ORF2 and ORF3 suggest that these three open reading
frames might be translationally coupled.

The deduced amino acid sequence of ORF] was analyzed by the Motif program
(Wisconsin Genetics Computer Group sequence analysis software) and shown to contain
a helix-turn-helix motif near the N-terminus (Figure 2.4), suggesting that the product of
ORF] is a DNA-binding protein (Brennan and Matthews, 1989; Harrison and Aggarwal,
1990). Analysis of the deduced amino acid sequence by the BLAST program supported

this observation by revealing that the ORF] product shared significant similarity to

62

Figure 2.4. Nucleotide sequence of 1.9-kb of DNA upstream of Tn5 lac Q4514. The
transcriptional start site is indicated by +1, and the -35 and -10 promoter regions are
marked. The complementary sequence of the primer used for the primer extension
analysis is; underlined. The putative translational start codons and potential ribosome
binding sites are boxed, and the deduced amino acid sequences of the Q4514 ORFs are
shown below the nucleotide sequence. The putative helix-tum-helix motif is shown in
bold letters. Restriction sites used to generate some deletions in the Q4514 upstream

region are also indicated.

63

71

141

211

281

351

421

491

561

631

701

771

841

911

981

1051

1121

1191

1261

1331

TGTACTGGCCCACCGCCTCCAACAGGGCCCGCATCTCCCGCCGGTGCTCCGCCAACGTGCCGGGCTGGGA
CATCAGCTCCCCGGCGAAGCGCAGGTCCTCCGCCACGTCCTCCAGCGTCTCCGCCACCCCGCGTGTGGCC
TCATCCAACTGGGCCTGGCGCTCCACCGCGAACTGGTGCCACTGCGCCTCCCGGTTGCGCTCCAACAGGG
TGAACACCCCCACCCCCACCGTCGTGAGGGCGACACAGAGCAACAGGATGGGGACGAGCGCGTACCGCAT
GGCCCGCGGAGTCTAACCGCCAGCCCCTCCCCAACCCCTTGAAATAGCAGGGAAATCAGGAGGCTCGCCG

SmaI -35 -10 1
GCCCGGGGGCCGACTTTGGCGATTGACACTCCGCGTCGGGCGCTGCTACCTACCGGTCEGTAGGTCTCdfl

Eco47III
GAAGGGACCAGA CGAACACCGGAGGACGGAAGCCGGACGAGGGCGAGCGCTACCGAGCCATCCTT
ORFl V T N T G G R K P D E G E R Y R A. I L

GAAACGGCGGCGCGGCTCATCTGTGACCGGGGGTACGAGGGCACGTCGATGCAGGAGATCGCCGCCGCGT
E T A. A R L I C D R G Y E G T S M Q E I A. A .A C

 

GCCGGATGACGAAGGCGGGCCTCTACCACCACATCCAGAACAAGGAGCAGTTGCTCTTCGCCATCATGAA
R.‘M T K A. G L Y H H I Q N K E Q L L F A I M N
NruI
CTACGGGATGGACCTGTTCGAGGAGCAGGTCCTCTCGCGGGTGCAGGACATCGCGAATCCGGTGGAGCGG
Y G M D L F E E Q V L S R V Q D I A N P V E R
SphI
CTTCGCGCGTGCATGCGCCACAACATCCTGCTGGTGACGCGGGGGTGGAGCAAGGAGGTCATCATCATCC
L R A C M R H N I L L V T R G W S K E V I I I L

TCCACGAGCACGCCACGCTCACCGGCGAGACGCGCGCCTTCATCGACGCCCGGAAGAAGAAGTACGTGGA
H E H A. T L T G E T R A F I D .A R K K K Y V D
StuI
CTTCCTGGAGGAGGCCTTTTCGCAGGCCTCGCAGCAGGGCCTCATCCGCCCCGTGGACCCGACGGTGGGC
F L E E A. F S Q A S Q Q G L I R P V D P T V G

GCCTTCTCGTTCCTGGGAATGGTGCTGTGGATCTACAAGTGGTTCAAGCCGGACGGGCGCCTCACGGATG
.A F S F L G M V L W I Y K W F K P D G R L T D E

AGCAGATCGCCGACGGCATGGTGGGCATGTTGTTCCCGCCCTTCGCCGCCGCTGGGGACACCGCTGGACA
Q I A D G M V G M L F P P F A .A .A G D T A G Q

GGCAGGGCCCTCCCCGTTGCGCATGGTGCCGAGCGTGTCGGCCACCGGCACGGATT-ACGC@A
AGPSPLRMVPSVSATGTDSEDA"

ORF2 M K
AGACGGCGCGCTGGTGCTCGCTGGAGGAGGCGGTGGCTTCCATTCCGGATGGCGCGTCGCTGGCCACCGG
T A. R W C S L E E A V .A S I P D G A. S L A T G

CGGTTTCATGCTGGGTCGCGCCCCCATGGCGCTGGTGATGGAGCTCATCGCGCAGGGCAAGCGCGACCTG
G F M L G R A P M A L V M E L I A Q G K R D L

GGCCTCATCTCCCTCCCCAACCCGCTGCCCGCGGAGTTCCTCGTGGCGGGCGGCTGTCTGGCCAGGCTGG
G L I S L P N P L P A E F L V A G G C L A R L E

AGATTGCCTTCGGCGCGCTGAGCCTCCAGGGCCGCGTGCGTCCCATGCCCTGCCTCAAGCGGGCCATGGA
I A F G A. L S L Q G R V R P M P C L K R A M E

64

1401

1471

1541

1611

1681

1751

1821

1891

GCAAGGCACCCTCGCCTGGCGCGAACATGATGGCTACCGCGTCGTCCAGCGGCTGCGCGCCGCGTCCATG
Q G T L .A W R E H D G Y R V V Q R L R A A S M

GGGCTGCCCTTCATCCCCGCGCCGGACGCGGACGTGTCCGGGCTGGCACGGACGGAGCCGCCTCCCACGG
G L P F I P A. P D A. D V S G L A. R T E P P P T V

TGGAGGACCCCTTCACCGGCCTGCGCGTGGCGGTGGAGCCTGCCTTCTATCCGGACGTGGCGTTGCTCCA
E D P F T G L R V A V E P A F Y P D V .A L L H

CGCGCGCGCCGCGGACGAGCGCGGCAACCTCTACATGGAAGACCCGACCACGGACCTGCTGGTGGCGGGC
.A R A A. D E R G N L Y M E D P T T D L L V A G

GCGGCGAAGCGGGTGATTGCCACGGTGGAGGAGCGGGTGGCGAAGCTGCCTCGCGCCACCCTGCCCGGCT
A A K R V I A T V E E R V A K L P R A T L P G F

TCCAGGTGGACCGCATCGTCCTGGCTCCCGGCGGCGCCCTGCCCAGCTGCGCCGGACTCTACCCGCACGA
Q V D R I V L A P G G .A L P S C .A G L Y P H D

CGACGAAATGCTGGCCCGCTACCTGTCGCTGGCGGAGACGGGCCGTGAAGCGGAGTTCCTGGAAACGTTG
D E M L A R Y L S L A E T G R E A. E F L E T L

CTGACG-CGGC@GCG

L T R R A A *
ORF3 M S

65

several transcriptional regulatory proteins. Among these is a putative transcription factor
of the TetR family identified in the Deinococcus radiodurans genome sequencing
project, to which ORFl exhibits 31% identity and 46% similarity over a 182 amino acid
stretch (White et al., 1999). ORF] also shows significant similarity to other TetR family
members. TetR is a repressor that regulates tetracycline resistance of many Gram-
negative bacteria (Hillen and Berens, 1994). In the absence of tetracycline, TetR binds to
operator sites, preventing transcription of tetA, the tetracycline resistance gene, and
transcription of tetR itself. When tetracycline is present, TetR binds to tetracycline and
releases the operators, so tetA and tetR are transcribed. These analyses suggest that the
protein product of the first open reading frame in the developmentally regulated Q4514
transcriptional unit might fiinction as a transcriptional regulator.

A BLAST search with ORF2 revealed significant similarity to several glutaconate
CoA transferases. The deduced amino acid sequence of ORF2 shows 29% identity and
44% similarity over its entire length to glutaconate CoA-transferase subunit A of
Acidaminococcusfermentans (Mack etal, 1994; Jacob etal, 1997). Glutaconate CoA
transferase is involved in glutamate fermentation. These results suggest that the Q4514
operon might encode components of an alternative metabolic pathway induced during M

xanthus development.

Precise localization of the Q4514 mRNA 5' end. The DNA sequence of the
Q4514 upstream region and the S1 nuclease protection assay results facilitated the design
of a primer for precise mapping of the location of the Q4514 mRNA 5' end by primer

extension analysis. The position of the primer is shown in Figure 2.4; however, the actual

primer contains a sequence complementary to that underlined. When primer extension

66

analysis was performed using RNA from DK1622 cells that had undergone 24 h of
development, a single primer extension product was identified (Figure 2.5). This localizes
the Q4514 mRNA 5' end to a guanine nucleotide at position 409 in the sequence (Figures
2.4 and 2.5). The mRNA 5’ end is 272 bp upstream of the Nrul site that was labeled in
the S1 nuclease protection experiment (Figure 2.3), so the results of the two methods are
in good agreement. No primer extension product was generated when mRNA prepared
from growing M xanthus cells was subjected to primer extension analysis (Figure 2.5).
Further deletion analysis of the Q4514 upstream region. The 1.62-kb DNA
segment upstream of Tn5 lac Q4514 extends to a SmaI site located at 54 bp upstream of
the putative transcriptional start site (Figures 2.1 and 2.4) and exhibits developmental
promoter activity (Figures 2.1 and 2.2). A 3' deletion was constructed to further define
the sequence required for developmental promoter activity. An Ec047III site (Figures 2.1
and 2.4) allowed construction of a 3' deletion to 65 bp downstream of the putative
transcriptional start site. The SmaI-Ec047III fragment was inserted into pREGl727 and
M xanthus strains with a single copy of the plasmid integrated at the chromosomal attB
site were tested for developmental promoter activity. Figure 2.6 shows that [3-
galactosidase activity reached a similar maximal level as for strains containing the 1.62-
kb Q4514 upstream segment (with its 5’ end also at -54 but its 3' end downstream of orj2)
fUSCd to lacZ, although there was a reproducible difference in lacZ expression at 6 h of
development (see discussion). These results demonstrate that a developmental promoter
exists between -54 bp and +65 bp relative to the location in the DNA of the Q4514

mRNA 5' end.

67

Figure 2.5. Primer extension analysis of Q45 14 mRNA. RNA was isolated fi'om wild-
type cells grown vegetatively (lane V) or cells that had undergone 24 h of development
(lane D). The same primer was used to sequence Q4514 upstream DNA. A portion of the
DNA sequence is indicated at the left. The arrow indicates the extension product that was
observed with RNA isolated from 24 h developing cells but not with RNA from
vegetatively growing cells. The asterisk marks the putative transcriptional start site as

determined by the position of the primer extension product.

68

ACGT V D

 

TCAGCCATCCAS
EGTCGGTAGGTT
*

69

Figure 2.6. Developmental expression of lacZ from a small segment containing the
Q4514 promoter. Developmental B-galactosidase specific activity was measured as
described previously (Kroos et al., 1986) for at least three independent transductants of
DK1622 containing a single copy of the 1.62-kb Q4514 upstream segment (0) or the
SmaI-Ec047III fragment (A) fiised to promoterless lacZ within pREGl727 and
integrated at Mx8 attB. The B-galactosidase specific activity of DK1622 containing the
pREGl727 vector alone with no insert integrated at Mx8 attB (I) is also shown. B-

galactosidase specific activity is expressed in nanomoles of o-nitrophenyl phosphate per T-

minute per milligram of protein.

 

[j

70

P

- n n b P

O 0 0 O O 0
6 5 4 3 2 1

33:00 050on ommEonflmmé

 

 

24 so 36 42 48
Time (Hr)

18

12

71

Q4514 can be transcribed by RNA polymerase isolated from growing M.
xanthus cells. Inspection of the DNA sequence upstream of the apparent Q4514
transcriptional start site revealed a perfect match in the —35 region to the E. coli 670
consensus (TTGACA), and a 2 out of 6 match to the —10 region consensus (TATAAT)
separated by 18 base pairs (Figure 2.4). To test whether RNA polymerase isolated from
growing M xanthus cells can transcribe from the Q4514 promoter, in vitro transcription
experiments were performed. Two different DNA segments containing the Q4514
promoter were used and run-off transcription products of the expected sizes were
observed (Figure 2.7A). M xanthus core RNA polymerase alone did not transcribe from
the Q4514 promoter (Figure 2.7B lane 1). Only when the core RNA pOlymerase was
mixed with 0A, the major vegetative 0 factor of M xanthus, was transcription observed
(Figure 2.7B lane 2). Primer extension analysis of RNA produced in vitro using
reconstituted 0" RNA polymerase mapped the 5’ end to the same position (data not
shown) as Q4514 mRNA produced in vivo (Figure 2.5). Together, these results indicate
that Q4514 can be transcribed by the major vegetative RNA polymerase of M xanthus,
suggesting that 0" RNA polymerase may be responsible for Q4514 transcription in vivo.

Since Q4514 is transcribed primarily, if not exclusively, during development
(Figures 2.2, 2.3, 2.5, and 2.6), we next ask whether oAis present during development.
Antiserum against B. subtilis 043 was used to detect (5“ in growing and developing M
xanthus cells (Figure 2.8) using Western blot analysis as described previously (Biran and

Kroos, 1997). The level of 0A remained fairly constant until 12 h into development, then

72

Figure 2.7. In vitro transcription from the Q4514 promoter. DNA fragments

containing the Q4514 promoter were transcribed in vitro with M. xanthus 0‘“ RNA
polymerase and the reactions were subjected to 5% polyacrylamide-8 M urea gel
electrophoresis. (A) Two different linear DNA templates containing the Q4514 promoter
region (SmaI-StuI and SmaI-Sphl fragments in lanes 1 and 2, respectively) were
transcribed with partially purified GA RNAP (Biran and Kroos, 1997). Run-off
transcription products of 458 (lane 1) and 303 bases (lane 2) are expected based on the
Q4514 mRNA 5’ end mapped in Figure 5. Arrows mark the expected run-off
transcription products. The upper band in each lane is the end-to-end transcript of the
template. The numbers on the left indicate lengths (in bases) of end-labeled MspI-
digested pBR3 22 marker fiagments. (B) Transcription from the SmaI-Sphl Q4514
upstream fragment by M xanthus core RNA polymerase alone (lane 1) or core RNA
polymerase plus 0A (lane 2). The arrOw marks the expected 303 base run-off transcription

product.

73

 

74

Figure 2.8. Western blot analysis of GA in growing and developing M xanthus cells.
DK1622 cells were collected at 0, 9, 12, 15, 18 and 24 h after the onset of development
(lanes 1 to 6), resuspended in sample buffer, and boiled for 5 min. Equal amounts of M
xanthus protein, as judged by measuring total protein in sonic lysates of the same samples,
were separated on 10% SDS-PAGE and electrotransferred to a nitrocellulose filter, which
was incubated with a polyclonal antiserum to B. subtilis 043 and visualized with goat anti-
rabbit IgG-horseradish peroxidase. The 0A is indicated by an arrow and protein standards

are marked in kDa.

75

 

\‘was'x‘e . Aimee. .

76

decreased markedly by 15 h. Thus, GA is present early in development when the level of
Q4514 mRNA rises (Figure 2.3).

Effect of an insertional mutation in ORF] on Q4514 expression. Since the
Q4514 promoter can be transcribed by GA RNA polymerase (Figure 2.7) and since 0" is
present in growing cells (Figure 2.8), we reasoned that Q4514 transcription must be
negatively regulated during growth, because Q4514 mRNA was not detected in growing
cells (Figures 2.3 and 2.5) and lacZ expression from Tn5 lac Q4514 was at a low
background level (Figures 2.2 and 2.6). The first ORF of the Q4514 operon is predicted
to encode a protein with a DNA-binding motif and sequence similarity to an auto-
repressor. Therefore, we hypothesized that the ORF 1 product functions as a repressor
which prevents Q4514 from being transcribed during growth. To test this hypothesis, an
insertional disruption mutation of 0rf1 was constructed. An Ec047III-Sphl restriction
fragment (Figure 2.1) from position 473 to 713 (Figure 2.4), which contains the putative
helix-turn-helix motif, was subcloned into an integrational vector and transformed into
wild-type DK1622. Transforrnants containing the plasmid integrated by homologous
recombination were identified by Southern blot hybridization (data not shown). The
effect of this orfI disruption mutation on Q4514 expression was tested by transforming
the mutant (MGVOI) with the plasmid (pTH10-3) containing the Q4514 promoter region
from -54 bp to +65 bp fiised to lacZ. Transforrnants containing a single copy of pTH10-3
integrated at Mx8 attB were identified by Southern blot hybridization (data not shown).
At least three of these transforrnants (MGV02) were assayed for B-galactosidase activity
during growth and development along with MTH10-3, the strain with a wild-type orfI

gene and a single copy of pTH10-3 at 0118. Figure 2.9 shows that significantly higher

77

Figure 2.9. Effect of 0rf1 and or]? mutations on developmental lacZ expression under
the control of the Q4514 promoter. Developmental B-galactosidase specific activity from
pTH10-3 integrated at 0118 was measured as described previously (Kroos et al., 1986) for

at least three independent isolates of M xanthus containing wild-type on"! and 0112 (A),
the 01f] mutation (O ), or the 0V? mutation (I). B—galactosidase specific activity is

expressed in nanomoles of 0-nitrophenyl phosphate per minute per milligram of protein.

78

 

 

/

 

180 -

— b p h p P h
0 0 0 0 0 0 0 O
6 4 2 0 8 6 4 2

33:00 050on 0005020555

 

0

18 24 30 36 42 48
Time (Hr)

12

79

lacZ expression was observed from the Q4514 promoter in the orfl mutant than in the
wild-type strain during growth and development.

To determine whether the higher lacZ expression observed in the 0111 mutant was
due to disruption of 0111, or to a polar effect on expression of 0112, I constructed an 0112
insertional disruption mutant using a similar approach. A DNA segment containing part
of 0rf2 from position 1164 to 1487 (Figure 2.4) was amplified by the PCR, verified by
DNA sequencing, and cloned into pBR322 to construct pTHl 1-1. This plasmid was
transformed into MTH10-3, the strain containing the Q4514 promoter firsed to lacZ at the
attB site in the chromosome. Transformants (MTHl 1-1) in which pTHl l-l had
integrated by homologous recombination, disrupting 0112, were identified by diagnostic
PCR (data not shown). As shown in Figure 2.9, the 0r12 mutation did not affect lacZ
expression from the Q4514 promoter. We conclude that disruption of orfl increases
Q4514 expression during growth and development. This conclusion is consistent with the
idea that ORF] protein represses Q4514 transcription directly, but we cannot rule out the
possibility that ORF] affects expression of lacZ fused to the Q4514 promoter indirectly.

Production of ORFl in E. coli and testing for binding to the Q4514 promoter.
To test whether ORFl protein binds to the Q4514 promoter region, I engineered E. coli
to overproduce ORF] or ORFl chimeras tagged with 6xHis at the N- or C-terminus;
using an IPTG-inducible T7 RNA polymerase expression system. As shown in Figure
2.10, a polypeptide of approximately the expected size (26 kDa) for ORFl-6xI-Iis (tagged
at the C-terminus) was enriched in cells after IPTG induction as compared to the
uninduced cells. E. coli overproduced ORFl or 6xHis-ORF1 (tagged at the N-terrninus)

upon IPTG induction (data not shown) about as well as ORF1—6xHis was overproduced

80

Figure 2.10. Production of ORF1-6xHis in E. coli and aflinity purification. Whole-cell
extracts (5 ul) of BL21(}.DE3) containing pTH191 designed to produce ORF1-6xHis
before (lane 1) and 2 or 3 h after IPTG induction (lanes 2 and 3). Eluted fraction (10 111)

from Ni-aflinity column (lane 4).

81

 

82

(Figure 2.10, lanes 2 and 3). The ORFl-6xHis protein was purified under nondenaturing
conditions with the aid of a Ni-aflinity column (Figure 2.10, lane 4).

To test whether affinity-purified ORF1-6xI-Iis binds to the Q4514 promoter,
mobility shift experiments were performed using a fragment, which spanned from -54 to
+65 bp, because expression from this segment was elevated in the 0111 mutant (Figure
2.9). Binding reactions incubated under standard conditions (Chodosh, 1988) or with
several modifications, such as changing the monovalent and divalent salt concentrations,
showed no specific shifted complex (data not shown). Likewise, no specific shifted
complex was observed with gel-purified ORF1-6xHis, ORFl, or 6xHis-ORF], or with
crude extracts from E. coli overproducing any of these proteins (data not shown). Also no
specific shifted complex was observed with a crude extract of growing M xanthus cells
or with a mixture of affinity-purified ORF1-6xHis and a crude extract of growing M
xanthus cells (data not shown). Thus, I did not observe specific binding of ORF]
products to the Q4514 promoter region under any of the conditions I tested

ORFl protein levels during M. xanthus growth and development. To measure
the ORFl level in M xanthus, the affinity-purified ORF1-6xHis fiision protein was used
to raise polyclonal antibodies in a rabbit. These anti-ORF1-6xI-Iis antibodies detected as
little as 0.3 ng of native ORFl overproduced in E. coli when diluted 1:5000 and used in
Western blot analysis (data not shown). However, the antiserum (as well as the
preirnmune serum) cross-reacted with many proteins in Western blot analysis of proteins
from growing M xanthus cells (data not shown). Therefore, the anti-ORF1-6xHis
antibodies were affinity-purified as described in Materials and Methods. Figure 2.11

shows that the affinity-purified antibodies detected a polypeptide in the extract of wild-

83

Figure 2.11. Detection of ORFl protein in extracts of growing M xanthus. Western blot
analysis of E. coli BL21 (ADE3) overproducing ORFl (lane 1), wild-type M xanthus
DK1622 (lane 2), and the M xanthus 0111 mutant was performed using affinity-purified
anti-ORF1-6xHis antibodies as described in Materials and Methods. The arrow indicates

ORF] protein and the positions of protein markers are indicated in kDa.

84

 

85

type M xanthus that was not present in the 0111 mutant extract. This polypeptide
migrated at a similar position as ORF] made in E. coli.

To determine whether the ORFl protein level changes during M xanthus
development, Tn5 lac Q4514-containing strain DK4514 was induced to develop and
samples were subjected to Western blot analysis and assayed for B-galactosidase specific
activity. Figure 2.12A shows that the ORF 1 protein level decreased slightly at 3-6 h then
rose later in development to a level higher than in growing cells. Figure 2.12B shows that
B-galactosidase activity increased slightly at 3-6 h, remained about the same at 9-12 h,
then rose later in development. Since ORFl negatively regulates Q4514 expression
directly or indirectly (Figure 2.9), it is possible that the slight decrease in the ORFl level
early in development allows the initial slight increase in Q4514 expression. However, the
major increase in Q4514 expression later in development does not correlate with a loss of
ORFl. Rather, the ORF] level increases, perhaps as a consequence of increased Q4514
promoter activity. The continued presence of ORF 1 during development is consistent
with the observation that the orfI mutant overexpresses Q4514 during development [”—
(Figure 2.9), suggesting that ORFl continues to negatively autoregulate Q4514
expression during development. Hence, the major increase in Q4514 expression later in

development appears to involve a mechanism other than loss or inactivation of ORFl.

 

Sporulation efficiency of Q4514 mutants. Tn5 lac Q4514 appears to be inserted at the
beginning of the third open reading frame of an operon. A previous study showed that
Tn5 lac Q4514 did not cause a developmental defect, as judged qualitatively by the

normal appearance of fruiting bodies (Kroos et al., 1986). To be more quantitative, and to

test whether the 0111 or 0112 insertional mutants exhibit development defects, the

86

Figure 2.12. Level of ORFl protein and B-galactosidase specific activity during
development of DK45 14. (A) Western blot analysis of proteins in extracts of M xanthus
DK4514 collected during growth (lane 2) and at 3, 6, 9, 12, 15, 18, 24 h after the onset of
development (lanes 3 to 9, respectively) using affinity-purified anti-ORFl-6xHis
antibodies as described in Materials and Methods. ORFl protein overproduced in E. coli
was detected in lane 1. Equal amounts of M xanthus protein, as judged by measuring
total protein in sonic lysates of the same samples, were subjected to SDS-PAGE and
Western blot analysis. The arrow indicates ORF] protein and the positions of protein
markers are indicated in kDa. (B) B-galactosidase specific activity of DK45 14 harvested

in the same experiment.

87

 

A

MW 036912151824
29—j “

     
    
     

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1234 5678 9
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120
$13,100
8.680
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o‘‘5 60
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(UP: 40
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0 3 6 9 12 15 18 21 24

Time (Hr)

88

sporulation efficiency was measured. The Tn5 lac Q4514-containing strain DK4514 had
a similar sporulation efficiency as wild-type DK1622 (Figure 2.13), indicating that
interruption of 0113 did not affect sporulation. The orfl and 0112 mutants appeared to
aggregate like wild type, but exhibited about fourfold and twofold reduced sporulation
efficiency, respectively (Figure 2.13), indicating that the 0111 and orf2 products play a

role in development.

89

Figure 2.13. Sporulation of Q45 14 mutants. Sporulation tests were performed as
described in Materials and Methods. The chart shows the average spore count of three

independent isolates of each strain and one standard deviation of the data.

90

Sporulation (1x10 5/ml)

140

120

100

(I)
O

O)
O

 

 

 

 

 

 

 

 

  

     

. x: ‘53-:- 1 . "31:“ . 1. ' .
“a . -. . . . ' -
1

wild type 0n‘1 0er Tn5 lac
mutant mutant Q4514

91

 

Discussion

We have cloned the DNA upstream of Tn5 lac Q4514 and identified a promoter
that is induced during M xanthus development. Promoter activity was lost in a graded
fashion in a 5’ deletion series, suggesting that multiple upstream elements contribute to
activity, but a deletion to —54 bp still permitted a low level of developmental induction.
Interestingly, the promoter shares sequence similarity with promoters of genes that are
transcribed during growth, and (IA RNA polymerase (RNAP), the major form of RNAP in
growing M xanthus, initiated transcription from this promoter in vitro. Our results show
that the Q4514 promoter is negatively regulated by ORF], the product of a downstream
gene. This negative autoregulation keeps Q4514 expression low during growth and early
in development. Full induction of the Q4514 operon during development involves
positive regulation acting through DNA elements both near the promoter and far
upstream.

Our deletion analysis of the Q4514 promoter region indicates that multiple DNA
elements spanning at least 650 bp upstream of the transcriptional start site contribute to
developmental promoter activity (Figures 2.1 and 2.2). Deletion of DNA between XhoI
and Sall sites located approximately 2.7 kb and 650 bp, respectively, upstream of the start
site, reduced developmental lacZ expression by about 30% (Figures 2.1 and 2.2).
Therefore, DNA more than 650 bp upstream of the start site is necessary for fiill Q4514
promoter activity. Likewise, deletion of DNA between the Sall site located
approximately 650 bp upstream of the start site, and the SmaI site located at -54 bp,

reduced developmental lacZ expression an additional 20%, indicating the importance of

92

this region (Figures 2.1 and 2.2). The graded loss of promoter activity for these deletions
was not specific to placement at the Mx8 attB site. Similar results were observed when
plasmids were integrated by homologous recombination upstream of Tn5 lac Q4514 to
create the corresponding 5' deletions firsed to lacZ at the native chromosomal location
(data not shown). The Q4514 promoter is not unique among developmentally regulated
M xanthus genes in requiring multiple upstream elements (Downard and Kroos, 1993).
The fps, csgA, and Q4499 promoters also exhibit large upstream regulatory regions (Kil
et al., 1990; Li et al., 1992; Fisseha et al., 1999). In the case of tps, multiple elements
appear to act over distances of several kb from the transcriptional start site (Kil et al.,
1990). In the cases of csgA and Q4499, upstream regions spanning more than 500 bp
were required for full expression, and in both cases expression was lost in a graded
fashion as 5' deletion endpoints approached the transcriptional start site (Li et al., 1992;
Fisseha et al., 1999). It was proposed that multiple transcription factors bind to these
upstream regions and regulate expression in response to different developmental cues.
Similarly, expression of the Q4514 promoter may involve several transcription factors [-
that respond to developmental cues by binding to upstream DNA elements and activating
transcription.

One cue that the Q4514 promoter does not respond to is C signaling. Intercellular

 

C signaling is mediated by the product of csgA (Shimkets et al., 1983; Shimkets and
Asher, 1988) and is required for the expression of many M xanthus genes that begin to
be expressed after 6 h into development (Kroos and Kaiser, 1987). Introduction of a csgA
mutation into cells containing Tn5 lac Q4514 delayed developmental lacZ expression

slightly in a previous study (Kroos and Kaiser, 1987). However, when we repeated this

93

experiment, we found no significant difference between wild-type and a csgA mutant
when we compared developmental expression from Tn5 lac Q4514. Also, expression of
5' deletions (i.e., pTH1-6, pTH4-l, and pTH4-3; see Figure 2.1 and Table 2.1) was tested
at Mx8 attB in a csgA mutant and was indistinguishable from that in wild-type cells (data
not shown). Moreover, when extracellular C signal was provided (by co-development
with wild-type cells) to the csgA mutants harboring the 5’ deletions, no increase in
expression was observed (data not shown), in contrast to the results observed for C
signal-dependent promoters (Fisseha et al., 1996; Brandner and Kroos, 1998; Fisseha et
al., 1999). Promoters that depend on C signaling for expression exhibit one or more
sequences (near —50 bp) matching the consensus CAYYCCY (Y means pyrimidine),
which has been called the C box (Fisseha et al., 1999). It has been speculated that C box
sequences are cis-acting regulatory elements important for the expression of C signal-
dependent genes. The Q4514 promoter does not have a sequence matching the C box
consensus near —50 bp. However, there are five sequences in Figure 2.4 that match the C
box consensus. Two of these are in the upstream region, centered at —191 (position 218 in
Figure 2.4) and -185 (position 224). These may be in the coding region of an upstream
gene, because an ORF that exhibits codon preference typical of M xanthus genes stops at

-113 (position 296) and extends upstream to the end of the region that has been

 

sequenced. The C-terminal polypeptide predicted by this partial ORF does not show
significant similarity to any protein in the databases. The other three sequences that
match the C box consensus are located in ORFl (centered at position 744 and 928 on the
transcribed strand) or ORF 2 (centered at position 1485 on the nontranscribed strand),

well downstream of the Q4514 promoter. A sequence matching the C box consensus is

94

expected to occur on average every 1200 bp in a high GC (70%) organism like M
xanthus, so the occurrence of C box sequences is higher than expected in the Q4514
region; but it remains to be tested whether any of these sequences are positioned properly
to influence Q4514 expression.

The Q4514 promoter is the first example of a developmental M xanthus promoter
that appears to be recognized by GA RNAP. Figure 2.14 compares the —35 and —10
regions of the Q4514 promoter with the corresponding regions of two promoters shown
previously to be transcribed by GA RNAP in vitro (Biran and Kroos, 1997), and with the
consensus -35 and —10 regions of promoters recognized by E. coli 070 and B. subtilis 0‘13
RNA polymerases. The Q4514 promoter has a perfect match in the —35 region to the
consensus. It resembles the vegA and ath promoters in that the --10 region is more GC-
rich than the consensus sequence. It was proposed previously that M xanthus 0A interacts

better with a more GC-rich —10 sequence than the primary sigma factors of E. coli and B.

subtillis (Biran and Kroos, 1997).

We showed that O’A persists at a high level at least until 12 h into development
(Figure 2.8), when the Q4514 mRNA level is also high (Figure 2.3). By 24 h into
development, 0" and Q4514 mRNA were barely detectable, yet B-galactosidase activity
from Tn5 lac Q4514 increased until 48 h of development (Figure 2.2). The continued rise
in lacZ expression after 18 h of development required upstream DNA beyond the SmaI
site located at —54 (Figures 2.2 and 2.6). We considered the possibility that another ‘
promoter located upstream of the SmaI site was expressed later in development, despite
detecting a single apparent mRNA 5' end (Figure 2.3). However, integration of a plasmid

(p1175SmSl4514, Table 2.1) into the chromosome (strain MGV03 in Table 2.1), placing

95

Figure 2.14. Comparison of promoter sequences. The —10 and —35 promoter regions and
the distance between them are shown for Q4514, for the M xanthus vegA (Komano et al.,
1987) and aphII (Rothstein and Reznikofl‘, 1981) genes, which are transcribed by GA
RNAP in vitro (Biran and Kroos, 1997), and for genes transcribed by E. coli 670 (Lisser
and Margalit, 1993) and B. subtilis 0'43 (Helmann, 1995) RNA polymerases. Matches to

70 / 0,43

the o consensus are indicated by boldface letters.

96

0451 4
vegA
aphll

670/643
consensus

-35
TTGACA
TAGACA

TTGCCA

TTGACA

97

18
17

17

17

-1 0
TACCTA
TAAGGG

TAAGGT

TATAAT

 

lacZ immediately downstream of the SmaI site, showed no developmental B-
galactosidase activity (Figure 2.1). Since there appears to be no additional promoter
upstream, we speculate that high-level expression of the Q4514-lacZ firsion mRN A at
12-18 h of development allows this message to persist later in development, giving rise to
continued B-galactosidase production. Another possibility is that Tn5 lac Q4514 may
disrupt or exert a polar effect on a gene whose product negatively regulates Q4514
expression. The level of Q45 14 mRNA was measured in wild-type cells in which the
postulated negative regulation would be intact, decreasing the mRN A level after 12-h
into development. In cells containing Tn5 lac Q4514, this negative autoregulatory loop
would be broken. Therefore, B-galactosidase activity would continue to increase later in
development. Both models predict that the level of Q45 14-lacZ fusion mRNA would
remain higher than was observed for the level of Q45 14 mRNA late in development.
The product of the first ORF downstream of the Q4514 promoter negatively
regulates expression of the Q4514 operon during both growth and development (Figure
2.9). Although ORF] has a helix-tum-helix DNA-binding motif and is most similar in
sequence to a family of repressors, we were unable to demonstrate specific binding of
ORF 1 to the Q4514 promoter region. It is possible that ORFl regulates another gene(s)
whose product(s) directly represses Q4514 transcription (Figure 2.15). Alternatively,
ORF] may be a direct repressor of Q4514 transcription, but may require an additional
factor that was not present under the conditions we tested. One such factor could be post-
translational modification of ORFl. However, we observed no significant difference in
migration on SDS-PAGE of ORFl produced in E. coli compared with ORF] produced

during M xanthus growth and development (Figures 2.11 and 2 12), providing no

98

 

evidence for post-translational modification of ORF 1. Perhaps ORF] requires binding of
a small molecule in order to bind specifically to the Q4514 promoter, analogous to the
case of tryptophan and the trp repressor (Joachimiak et al., 1983).

Developmental induction of Q45 14 expression can occur in the absence of ORF]
and in the absence of upstream DNA beyond ~54 (Figure 2.9). This suggests that 0A
RNAP transcribes from the Q4514 promoter more efl'rciently during development than
during growth, because the level of 0“ remains about the same during growth and early
in development (Figure 2.8). If many oA-dependent genes are repressed as cells enter
development, perhaps 0" RNAP becomes available to transcribe the Q4514 promoter.
Another possibility is that a transcriptional activator is induced upon starvation and binds
between ~54 and +65 in the Q4514 promoter region.

Insight into the regulation of Q45 14 expression can be gained fi'om comparison of
the level and timing of developmental lacZ expression observed for different promoter
constructs. B-galactosidase activity from the promoter region between ~54 and +65
reached 160 U in the absence of ORFl (Figure 2.9). However, B-galactosidase activity
reached nearly 400 U from the Q4514 promoter with additional upstream DNA, even
when ORFl was present (Figure 2.2). Clearly, upstream DNA elements enhance Q4514
promoter activity. It will be interesting to test the activity of the Q4514 promoter with
upstream elements in the absence of ORF]. We might observe a higher maximum level
of expression, as suggested by the results shown for the Q4514 promoter from ~54 to +65
in the presence or absence of orfl at the native chromosomal site (Figure 2.9). We might
also observe a change in the timing of lacZ expression. Figure 2.9 shows that lacZ

expression increased significantly by 4 h of development for strains with lacZ fused at

99

+65. This is most obvious for the strain completely lacking ORF] due to a disruption at
the native site (Figure 2.9, diamonds). The other strains examined in this experiment had
orfl intact at the native site, and therefore exhibited a lower level of expression due to
ORF] negative regulation in trans (since the lacZ fusion is located at the Mx8 attB site),
but like the strain completely lacking 0111, these strains showed no obvious lag in lacZ
expression early in development (Figure 2.9). One of these strains was also used in the
experiment shown in Figure 2.6 (triangles), and it is clear that expression increased by 6h
of development. In contrast, lacZ expression appeared to be delayed, increasing
dramatically only after 6h of development, for strains with lacZ fused downstream of 0112
(Figure 2.2 and Figure 2.6, diamonds). In these strains orfl is expressed from the Q4514
promoter at Mx8 attB as well as from the native site, so ORFl may be responsible for the
delay in expression, by directly or indirectly inhibiting Q4514 promoter activity (Figure
2.15). We cannot exclude the possibility that DNA between +65 and the site of Tn5 lac
Q4514 insertion (downstream of 0112) causes the delay in developmental expression, due
to regulation at the transcriptional or translational level.

If ORF] is responsible for delaying Q4514 expression early in development, how
is this negative regulation overcome? There was a small decrease in the level of ORF] at
3-6 h of development that correlated with a small increase in expression from Tn5 lac
Q4514, but the major rise in lacZ expression did not correlate with a loss of ORFl
(Figure 2.12). The finding that lacZ expression from the Q4514 promoter continues to
rise during the first 36 h of development in 0111 mutant cells, but not in cells expressing
orfl (Figure 2.9), suggests that ORFl continues to negatively regulate Q4514 expression

during development. We speculate that positive regulation by one or more transcriptional

100

Figure 2.15. A model describing regulation of the Q4514 operon. During grth and
development, ORF] negatively regulates transcription by (TA RNAP either by binding
directly to the promoter region or by regulating another gene(s) whose product(s) (x)
represses transcription. Developmental induction may initially involve a decrease in
ORFl-mediated negative regulation early in development but our data suggest that ORFl
continues to negatively autoregulate Q4514 expression during development and we
propose that a transcriptional activator(s) is primarily responsible for developmental

induction of Q4514.

101

6A RNAP

Transcriptional j

 

 

 

Activator(s)
\ 0rf1 . orf2 0113?
I l [1 r1—
17 \ l 1 site of
04514
? ORF1 ORF2 insertion

X I DNA-binding glutaconate
Protein GOA-transferase

102

activators overcomes ORF1-mediated negative regulation and causes the major rise in
Q4514 transcription during development (Figure 2.15).

Disruption of orfl caused a mild sporulation defect (Figure 2.13). This may be
due to a polar effect on expression of 0112 since disruption of 0112 also caused a mild
sporulation defect (Figure 2.13). ORF2 shows similarity to glutaconate CoA-transferase
subunit A of Acidaminococcus fermenians and related proteins in other organisms. These
enzymes catalyze transfer of the CoA moiety from acetyl-CoA to 2-hydroxyglutarate and
related compounds like glutamate in the glutamate fermentation pathway (Mack et al,
1994). Our results suggest that the Q4514 operon might encode components of an
alternative metabolic pathway that is induced during M xanthus development and plays a
role in sporulation. Sequence analysis suggested the possible existence of a third ORF
overlapping the ORF2 stop codon and interrupted after just two amino acids by Tn5 lac
Q4514. In Acidaminococcusfermentans, subunit B of glutaconate CoA-transferase is
encoded in the same operon downstream of subunit A. By analogy, ORF3 might encode
subunit B of glutaconate CoA-transferase. However, DK4514, in which 0113 is disrupted
by Tn5 lac Q4514, did not show a sporulation defect (Figure 2.13). It is unlikely that a
functional ORF3 protein is made in DK4514, because the Tn5 lac insertion disconnects
0rf3 from its normal transcription and translation start signals. Although many questions
remain about the fimction and regulation of the Q4514 operon, it is the first example of a
developmentally regulated M xanthus operon that is transcribed by the major vegetative
RNAP, and it is subject to complex control, including negative autoregulation by the
product of the first gene in the operon and positive regulation by upstream DNA

elements.

103

Chapter III

Identification of regulatory proteins involved in the
expression of a C-signal-dependent gene

104

Abstract

Q4403 is a Tn5 lac insertion in the Myxococcus xanthus chromosome that fiises lacZ
expression to a developmental promoter, which is expressed between 6 and 12 h after
starvation. Cell-cell interactions, including C-signaling, are required for the
developmental expression of Tn5 lac Q4403. It has been shown previously that DNA
downstream of ~80 bp relative to the Q4403 transcriptional start site is sufficient for C-
signal-dependent developmental expression of lacZ, comparable to that observed with
much larger upstream segments. However, the developmental lacZ expression was
abolished upon deletion to ~72 bp, suggesting the possible involvement of an upstream
activator protein that interacts with DNA between ~80 and ~72. To search for this
protein, I first cloned DNA from ~103 to +3 6, which has less sequence downstream of
the transcriptional start site than the previously tested constructs, and showed that it had
similar promoter activity. Therefore, this fragment was used to search for a regulatory
protein(s) that interacts with the Q4403 promoter region. I identified a development-
specific complex using an electrophoretic mobility shift assay. It appears that a protein
binds between ~41 and ~11, and the formation of the shifted complex does not depend on
C-signaling, or on A-, D-, or E-signaling. Shifted complexes were also formed with the
C-signal-dependent Q4400 promoter and with three promoters that are not C-signal-

dependent (Q4514, vegA and aphll). Taken together, the data suggest that the protein

forming the shifted complex with the Q4403 promoter fi'agment is unlikely to mediate C-

signaling-dependent expressidn of Q4403.

105

Introduction

Cell-cell interactions play a critical role in multicellular development and cellular
differentiation. Myxococcus xanthus is a gram-negative, rod-shaped soil bacterium that
undergoes multicellular development when nutrients are depleted (Dworkin and Kaiser,
1993). When cells are starved at a high density on a solid surface, they move in a
coordinated fashion into aggregation centers and form mound-shaped multicellular
structures known as fruiting bodies, inside of which most of the cells lyse and some cells
differentiated into ovoid spores. When nutrients are available, these dormant, heat- and
sonication-resistant spores germinate.

This multicellular process requires cell motility and communication between cells
(Shimkets, 1999). At least five extracellular signaling interactions have been identified
and are known as the A-, B-, C-, D-, and E-signaling interactions (Hagen et al., 1978;
LaRossa et al., 1983; Downard et al., 1993). Mutants unable to produce any one of these
signals are arrested in development, but the defects are rescued upon codevelopment with
wild-type cells or mutants of a different signaling group. Cells defective in different
signaling interactions arrest development at difl‘erent stages. Mutants defective in A-, B-,
D-, or E-signaling block development before 5 h, whereas C-signaling affects genes that
are expressed after 6 h into development (Downard et al., 1993; Kaiser and Kroos, 1993;
Downard and Toal, 1995).

Our studies have focused on the regulation of developmental gene expression by C-
signaling. The csg group of mutants, which are defective in C-signaling, are genetically

simple and well-characterized. All csg mutations map to a single locus in the

106

 

chromosome, csgA, and the cells are defective in rippling, aggregation, fruiting body
formation and sporulation (Hagen et al., 1978; Shimkets and Asher, 1988). Almost all
genes expressed later than 6 h into development exhibit reduced expression or no
expression in a csgA mutant background (Kroos and Kaiser, 1987). The csgA gene
appears to encode a 25-kDa protein of the short-chain alcohol dehydrogenase family and
the ngA protein is associated with the cell surface and/or the extracellular matrix
(Shimkets and Rafiee, 1990; Lee et al., 1995). Efficient C-signal transmission requires
cell movement (Kroos et al., 1988; Kim and Kaiser, 1990b). During development, cells
move into alignment and become densely packed in the outer domain of a nascent
fruiting body (Sager and Kaiser, 1993). It has been proposed that end-to-end contact
brought about by cell movement mediates efficient C-signaling (Sager and Kaiser, 1994).
C-signaling is self-reinforcing. ngA affects the gliding movements of starved individual
cells in a way that is thought to promote aggregation, thus bringing about cell alignment
that permits more efficient C-signaling (Sogaard-Anderson and Kaiser, 1996; Jelsbak and
Sogaard-Anderson, 1999). Also, ngA addition to starved cells stimulates expression of
csgA, creating a positive feedback loop (Kim and Kaiser, 1991). A lower level of C-
signaling is sufficient for aggregation and expression of genes partially dependent on C-
signaling, whereas a higher level of C-signaling is required for sporulation and
expression of genes absolutely dependent on C-signaling (Kim and Kaiser, 1991; Li et
al., 1992). Thus, ngA appears to act as a developmental timer that triggers the normal
developmental sequence of aggregation, fiuiting body formation and finally sporulation

as the ngA concentration increases during development.

107

Sixteen C-signal-dependent genes have been identified by using Tn5 lac, a
transposon that contains a promoterless lacZ (Kroos and Kaiser, 1984). Transposition of
Tn5 lac into the M xanthus chromosome can generate a transcriptional fusion between
lacZ and an upstream M xanthus promoter (Kroos et al., 1986). Tn5 lac insertions in C-
signal-dependent genes were identified by comparing lacZ expression in wild-type cells
and csgA mutants (Kroos and Kaiser, 1987).

The expression of Tn5 lac Q4403 is absolutely dependent on C-signaling (Kroos
and Kaiser, 1987). The Q4403 promoter has been localized and mapped (Fisseha et al.,
1996). It was shown that DNA downstream of ~80 is sufficient to direct C-signal-
dependent expression, whereas the expression is completely abolished when 8 bp more is
deleted from the 5’ end. Since DNA from ~80 to ~72 is upstream of the typical RNAP
binding site, we speculate that a regulatory protein(s) binds to this region and mediates C-
signal-dependent expression. To search for this protein and any other proteins that
interact with the Q4403 upstream region, I first cloned a small Q4403 promoter-
containing DNA segment (~103 to +3 6) and showed that this segment has similar
promoter activity as DNA segments with more downstream sequence. The small DNA
segment was used as a probe to search for DNA-binding proteins. A development-
specific, DNA-binding activity was identified. However, it does not require the ~80 to ~
72 sequence to bind, nor does binding require C-signaling. Moreover, the DNA-binding
activity did not appear to be specific for C-signal-dependent promoters, so it is unlikely

to be responsible for C-signal-dependent gene expression.

108

Materials and Methods

Bacterial strains and plasmids. Strains and plasmids used in this work are listed
in Table 3.1.

Growth and development. Escherichia coli cells were grown at 37°C in LB
medium (Sambrook et al., 1989) containing 50 pg ampicillin or 25 pg kanamycin per ml,
as necessary. M xanthus was grown at 32°C in CTT medium (Hodgkin and Kaiser, 1977)
in liquid culture or on agar (1.5%) plates with 40 pg kanamycin or 12.5 pg
oxytetracycline per ml when required (Kroos et al., 1986). Fruiting body development
was performed on TPM (10 mM Tris-HCl, pH 8.0, 1 mM KH2P04, 8 mM MgSOr, final
pH 7.6) agar (1.5%) plates as described previously (Kroos et al., 1986).

Molecular cloning. Recombinant DNA work was performed using standard
techniques (Sambrook et al., 1989). Plasmid DNA was prepared from E. coli DHSo. or
JMSB.

To construct a pREGl 727 derivative with a small DNA segment containing the
Tn5 lac Q4403 promoter, a DNA fragment from ~103 to +36 bp relative to the Q4403
transcriptional start site was generated by PCR using pMF3.4 as a template. The
upstream primer was 5’-CTCTCTAGAAGACATCGCGCCTTGAA-3’, which has an
underlined XbaI site and anneals to the sequence between positions ~103 and ~87. The
downstream primer was 5’-CTCGGATCCTTCATGTTTTACCCAGA-3’, which has an
underlined BamHI site and anneals to the sequence between position +20 and +36. The
amplified fragment was digested with BamHI and XbaI, gel-purified and ligated to
BamHI/XbaI-digested pREGl727 to construct pTH6-1. The insert was sequenced to

ensure that no errors occurred during the PCR.

109

Table 3.1. Bacterial Strains and Plasmids

 

 

 

Strain or plasmid Characteristics Source

E. coli

DHSa supE44 AlacUl69 (1)80 AlacZ AMIS hst17 Hanahan, 1983

recAl endAl gyrA thi-lrelAl

JM83 ara Alac-pro strA thi (bdlacZ AM15 Messing, 1979

M xanthus

DK1622 wild type Kaiser, 1979

MMF1727 atthszEGl727 Fisseha et al., 1996

MTH6-1 attB: :pTH6-1 This study

MMFIOO attB::pMF100 Fisseha et al., 1996

DK4746 Tn5 Q4678 asgB480 Dale Kaiser

DK5270 Tn5 lac (Km‘) Q4403 csgA : :Tn5-132 (Tc’) Q205 Kroos and Kaiser,
1987

DK3261 Tn5 Q1867 dsg439 Cheng and Kaiser, '7'“
1989b

ID300 esgzzTn5 (Q258) J. Downard

Plasmid

pUC19 Apr laca Yanisch-Perron et al.,
1985

pMF3.4 Apr (pUCl9)°, 674-bp SphI-BamI-II fragment Fisseha et al., 1996

upstream of Q4403 Tn5 lac
pMFlOO Apr Kmr (pREGl727), 521-bp HaeII-Baml-H Fisseha et al., 1996

fragment upstream of Q4403 Tn5 lac

110

pREGl727 Apr Kml' Pl-inc ath lacZ

pTH6-1 Apr Kmr (pREGl727), DNA from -103 to +36
bp of Q4403 transcription start site generated by
PCR using pMF3 .4

pTHlO-l Apr (pUCl9); 120-bp SmaI-Ec047III fragment
from pTH4-4

Fisseha et al., 1996

This study

Chapter 2

 

a The vector is indicated in parentheses.

111

Construction of M. xanthus strains. Strain MTH6-l was constructed by P1
specialized transduction of pTH6-l from the rec+ E. coli strain JM83 into wild-type
DK1622 (Gill et al., 1988; Fisseha et al., 1996). Because of the presense of attP on the
plasmid, pTH6-1 integrated into the M xanthus chromosome at Mx8 attB. At least three
derivatives, each containing a single copy of integrated plasmid, were identified by
Southern blot analysis (Sambrook et al., 1989; Fisseha et al., 1996), and B—galactosidase
production was measured under developmental conditions as described previously (Kroos
et al., 1986).

Crude extract preparation for gel mobility shift assay. Vlfrld-type DK1622
cells or signaling mutant cells were grown in CTT liquid medium with appropriate
selection until the density reached 100 Klett units. Cells were then sedimented at 43 00 x
g for 10 min. The cell pellet was resuspended to 2000 Klett units using TPM buffer and
0.5 m1 aliquots were spread on TPM plates and incubated at 32°C. Vegetative and
developmental M xanthus cells collected at the indicated time points were harvested and
stored at ~70°C. Extracts were prepared by the method of Gross and Burgess (Gross et
al., 1976) with some modifications. Thawed cell pellets (approximately 0.4 ml) were
suspended in 0.25 ml of solution A (10 mM Tris-HCl [pH 8.0], 25% sucrose, 100 mM
NaCl, 1 mM phenylmethylsulphonyl floride [PMSF]), followed by addition of a mixture
of protease inhibitors (around 5 pl total) of 1 pg aprotinin, 0.5 pg leupeptin and 0.7 pg
pepstatin A. The mixture was incubated on ice for 15 nrin. This was followed by addition
of 0.06 ml of solution B (300 mM Tris-HCl [pH 8.0], 100 mM EDTA, 4 mg/ml
lysozyme, 1 mM PMSF) and incubation on ice for 10 min. Finally, 0.31 ml of solution C

(l M NaCl, 20 mM EDTA, 0.08% deoxycholate, 1 mM PMSF) was added and the

1.12

mixture was incubated at 10°C for 10 min. The resulting lysates were centrifugated for 40
min at 30,000 x g. The protein concentration of the supematants was quantified by the
Bradford method (Bradford, 1976). The extracts were used immediately for mobility shift
assays or quick-frozen in a 95% ethanol/dry ice bath after addition of glycerol to 50%
final concentration.

Mobility shift experiments. PCR products containing Q4403 upstream DNAs
were 5' end-labeled with [y-32P] ATP and T4 polynucleotide kinase. A restriction
fragment from ~105 to +53 relative to the Q4403 transcriptional start site, obtained from
pMF3.4 digested with MscI and anI, was gel-purified, phosphatase-treated, and 5' end-
labeled with [7-321)] ATP and T4 polynucleotide kinase. The labeled DNAs were purified
with a QIAquick Nucleotide Removal Kit (Qiagen). Ten pg of crude extract was
incubated with 10,000 cpm of probe (typically 0.75 ng) and 1 pg of poly(dI-dC) as
nonspecific competitor at 30°C for 15 min to allow DNA binding. The 20 pl binding
reaction contained 12% glycerol, 12 mM HEPES (pH 7.9), 200 mM NaCl, 4 mM Tris-
HCl (pH 7 .9), 0.6 mM EDTA, and 5 mM MgC12. The samples were then subjected to

electrophoresis on 5% native polyacrylamide gels at 4°C in low ionic strength buffer

(Chodosh, 1988). After electrophoresis, the gels were dried for autoradiography.

113

Results

Further deletion analysis of the Q4403 promoter region. Q4403 is a Tn5 lac
fusion that is expressed only during development in a C-signaling dependent manner. A
previous study showed that DNA between ~80 and +382 bp relative to the transcriptional
start site was sufficient for C-signal-dependent promoter activity, but a deletion that
removed 8 bp from the upstream end of this fragment abolished promoter activity
(F isseha et al., 1996). To better define the downstream boundary of DNA needed for
promoter activity, DNA between ~103 and +36 bp was amplified by PCR and cloned into
pREGl727 upstream of a promoterless lacZ gene, resulting in pTH6-1. pTH6-1 was
integrated into the M xanthus chromosome at the myxophage Mx8 attB. Transductants
containing a single copy of pTH6-l integrated at attB were identified by Southern blot
hybridization (data not shown). Several of these transductants were assayed for B-
galactosidase activity during development and showed similar promoter activity as
compared to MMFIOO, a strain containing Q4403 DNA between ~80 and +3 82 bp fused
to lacZ at the attB site (Figure 3.1).

Detection of a development-specific shifted complex with the Q4403

promoter region. To search for a protein(s) that interacts with the Q4403 promoter
region, a PCR product containing DNA between ~103 and +36 bp or a restriction
fragment containing DNA between -105 and +53 bp was utilized in electrophoretic
mobility shift assays. Crude extracts from vegetatively growing or developing wild-type
M xanthus DK1622 were prepared and used in mobility shift assays as described in
Materials and Methods. A shifted complex was detected with extract made from cells

harvested 12 h into development, in the absence or the presence of the nonspecific

114

Figure 3.1. Developmental expression of lacZ from a DNA segment containing the
Q4403 promoter. Developmental B-galactosidase specific activity was measured as
described previously (Kroos et al., 1986) for at least three independent transductants of
DK1622 containing a single copy of the Q4403 upstream segment from ~80 to +3 82 bp
(4) or the DNA segment from ~103 to +36 bp relative to the Q4403 transcriptional start

site (I). B-galactosidase specific activity is expressed in nanomoles of o-nitrophenyl

phosphate per minute per milligram of protein.

115

Time (Hr)

 

r

 

35

33:00 050000 0005090335

116

competitor poly(dI-dC), whereas no shifted complex was detected with vegetative cell
extract (Figure 3.2).

Competition assays with unlabeled fragments. To test whether the formation of
the shifted complex is specific to the probe DNA, competition experiments including an
excess amount of the same unlabeled DNA in the binding reaction were performed in the
presence of the nonspecific competitor poly(dI-dC). The intensity of the shifted band
started to decrease with a 10x molar excess of the unlabeled DNA and became very faint
with a 100x molar excess of the unlabeled DNA (Figure 3.3). These results show that the
DNA-binding activity is somewhat specific for the Q4403 promoter containing fi'agment.

Developmental timing of formation of the shifted complex. To determine the
level of the DNA-binding activity during development, crude extracts made from
DK1622 cells harvested at 4, 8, 12, 20 or 24 h into development, as well as during
vegetative growth, were tested. A shifted complex was observed at 4 h into development,
reached its strongest intensity at 12 h, and disappeared after 24 h into development
(Figure 3 .4).

Dependence of the DNA-binding activity on extracellular signals. The
expression of Q4403 is absolutely dependent on C-signaling (Kroos and Kaiser, 1987).
To test whether the DNA-binding activity exhibits dependency on C-signaling, crude
extracts made from vegetative and developmental csgA mutant cells were tested. A
similar shifted complex was observed with developmental csgA mutant cells (Figure
35A), suggesting that neither the production nor the DNA-binding activity of the protein
in the shifted complex is dependent on C-signaling. No shifted complex was seen with

vegetative cell extracts.

117

Figure 3.2. Gel mobility shift analysis of the Q4403 promoter region. A PCR product
spanning ~103 to +36 bp was used as probe with extracts made from vegetatively
growing (T0) or 12-h developing (T12) wild-type DK1622 M xanthus cells in a mobility
shift experiment as described in Materials and Methods. Poly(dI-dC) was added to
binding reactions, unless indicated otherwise. The arrow indicates the position of a

shifted complex with the developmental extract.

 

118

T0 T12
poly(dl-dC) - + - +

 

 

 

119

Figure 3.3. Competition with unlabeled DNA. The binding reaction was carried out with
0.75 ng (7 finol) 3‘zP-labeled Q4403 DNA fragment (~105 to +53 bp) alone (lane 1) or
with the addition of extract (10 pg protein) made from 12 h-developing DK1622 cells
and lpg poly(dI-dC) (lanes 2-5). The same Q4403 DNA fragment, except unlabelled, was

also included in the binding reaction in lO-fold (lane3), 50-fold (lane 4) or lOO-fold (lane

5) molar excess over the labeled probe. The shifted complex is marked by the arrow.

120

competitor — 10x 50x 100x

 

121

Figure 3.4. Developmental timing of formation of the shifted complex. Extracts (10 pg
protein) made from DK1622 cells collected during growth (lane 2) or at 4,8, 12, 20 or 24
h after starvation (lanes 3 to 7) were incubated with 0.75 ng 32P-labeled Q4403 DNA
probe (-105 to +53 bp) and lpg poly(dI-dC) at 30°C for 15 min. The mixtures were then

subjected to gel mobility shift analysis. Lane 1 is the probe alone and the arrow indicates

the shifted complex.

122

 

123

The dependence of DNA-binding activity on three other extracellular signaling
interactions (A-, D- and E-signals) was also tested using extracts prepared from
signaling-defective mutant cells that had undergone development. The DNA-binding
activity was present in all three mutant extracts (Figure 3.5B). Together, these results
indicate that the DNA-binding activity is not dependent on the A-, C-, D-, or E-signaling
interactions.

Localization of DNA sequence in the Q4403 promoter region required for
binding. To localize the DNA sequence that was recognized by the DNA-binding
activity, a series of 5' deletions were generated by PCR. The deletions all have the same
3' end (at +3 6) and were used as labeled probes in binding reactions. As shown in Figure
3.6, a shifted complex was detected with DNA fragments with 5’ ends at -80, -72, or -41
bp in the absence or presence of 1 pg poly(dI-dC). When DNA with a 5’ end at -31 bp
was used as probe, a shifted complex with much weaker intensity was observed without
poly(dI-dC) and disappeared when 1 pg poly(dI-dC) was included in the reaction. This
result suggests that DNA between ~41 and ~31 contributes to formation of the shifted
complex.

These deletion segments, as well as DNAs from ~105 to ~11 and from -11 to +53
bp, were also used as competitors in experiments with DNA between -105 and +53 as
probe. The deletions with 5’ ends at -80, -72 or -41 bp (all having the same 3’ end at +3 6)
and the DNA from ~105 to ~11 competed away the formation of the shifted complex as
well as full length fragment (-105 to +53), whereas DNA from -11 to +53 bp failed to
compete (data not shown). Taken together, the results suggest that a protein binds to

Q4403 promoter DNA between -41 and -11 bp relative to the transcriptional start site.

124

Figure 3.5. Dependence of the DNA-binding activity on extracellular signals. (A)
Extracts (10 pg protein) from csgA cells collected during grth (lane 2) or at 4, 8,12, 20
or 24 h after starvation were incubated with 0.75 ng 32P-labeled Q4403 DNA probe (-105
to +53 bp) and 1 pg poly(dI-dC) at 30°C for 15 min. The mixtures were then subjected to
electrophoresis on a 5% native polyacrylamide gel. (B) Extracts (10 pg protein) made
from wild-type cells (lane 2) or asg (lane 3), dsg (lane 4), or esg (lane 5) cells harvested
at 12 h into development were incubated with probe as described for panel A. No extract

was added to probe in lane 1 of both panels. Shifted complexes are indicated by the I

arrows.

 

125

 

126

Figure 3.6. Gel mobility shift assays of 5' deletion fragments containing the Q4403
promoter. Q4403 DNA (40 fmol) from ~80 to +36, -72 to +36, -41 to +36, or ~31 to +36
was incubated with extract (10 pg protein) fi'om wild-type DK1622 cells at 12 h into
development in the presence or absence of 1 pg poly(dI-dC) at 30°C for 15 min. The

mixtures were then subjected to electrophoresis on a 5% native polyacrylamide gel.

127

probe -80~+36 -72~+36 -41~+36 -31~+36

poly(dldC)--+--+--+--+
extract-++-++_++-++

 

 

 

128

Binding to other promoters. Do the extracts fiom developing cells possess
DNA-binding activity for other promoters, or is the Q4403 promoter unique in this
respect? To answer this question, two other M xanthus developmental promoter-
containing DNAs were tested. Q4514 DNA spanning fiom -54 to +65 bp (chapter 2) was
shifted in a similar fashion as Q4403 promoter DNA, both in terms of the amount of
shifted complex produced and its migration (data not shown). Likewise, Q4400 DNA
spanning ~101 to +101 bp (Brandner and Kroos, 1998) was shifted similarly (data not
shown). Hence, the extract shifts C-signal-dependent (Q4403 and Q4400) and C-signal-
independent (Q4514) developmental promoters. I also tested two M xanthus vegetative
promoter DNAs; vegA, spanning from ~3 20 to +100 bp and aphll, spanning from ~250 to
+70 bp. A similar shifted complex was detected with vegA DNA (Figure 3.7). To ask
whether the same protein is binding to vegA and Q4403 promoter DNAs, a competition
experiment was performed. A 50-fold molar excess of vegA DNA competed away the
shifted complex formed with Q4403 promoter DNA (data not shown); suggesting that
same protein binds to both promoters. A more abundant and perhaps slightly slower-
rnigrating complex was detected with aphll DNA (Figure 3.7). No shifted complex was
detected when part of the coding sequence of Q45 14 DNA (from +5 to +273 bp relative
to the transcriptional start site) was tested (data not shown). This shows that not every
DNA segment can be shifted by proteins in developing cell extracts and suggests that

complex formation may require promoter DNA.

129

Figure 3.7. Gel shift experiments with different promoters. Q4403 DNA (~103 to +36
bp), vegA DNA (—3 20 to +100 bp) or aphll DNA (~250 to +70 bp) was incubated with
extract (10 pg protein) of DK1622 cells 12 h into development in the presence or absence
of 1 pg poly(dI-dC) at 30°C for 15 min. The mixtures were then subjected to

eletrophoresis on a 5% native polyacrylamide gel.

130

probe Q4403 vegA aphll
poly(dl-dC) - + - - - + - - +
extract - + - - .+ + - + +

 

 

131

Discussion

A previous study showed that elements needed for C-signal-dependent expression
of Q4403 lie between ~80 and +3 82 bp relative to the transcriptional start site (F iseha et
al., 1996). In this study, I showed that Q4403 DNA between ~103 and +36 bp has similar
developmental promoter activity as the ~80 to +3 82 bp segment (Figure 3.1). Together,
the results localize the Q4403 promoter between ~80 and +36 bp.

The principal aim of this study was to identify a regulatory protein that binds to
the Q4403 upstream region and activates transcription in response to C-signaling. DNA
between ~80 and ~72 bp is critical for Q4403 promoter activity, since deletion of this ,
sequence abolishes developmental expression (Fisseha et al., 1996). Initially, a synthetic,
double-stranded DNA with sequence identical to ~92 to ~64 bp of the Q4403 promoter
was used as a probe in gel mobility shift experiments to search for a DNA-binding
protein. No specific shifted complex was obtained using the conditions described in
Materials and Methods, or using a variety of modifications (data not shown). We
reasoned that the synthetic DNA might be too short to allow binding. Therefore, larger
Q4403 promoter DNA fragments were used as probes in mobility shift experiments.
Using DNA between ~103 and +3 6, a development-specific shifted complex was
observed (Figure 3 .2). A competition experiment with unlabelled Q4403 promoter DNA
demonstrated that binding was somewhat specific for the Q4403 promoter (Figure 3.3). A

10-fold molar excess of specific competitor was sufficient to reduce formation of the
shifted complex in the presence of more than 1300-fold molar excess of nonspecific

competitor. Because the crude extracts used in my experiments contain many DNA-

132

binding proteins, some of which may bind to the Q4403 promoter region with some
specificity, it may not be surprising that a 100-fold molar excess of specific competitor
was needed to virtually eliminate formation of the shifted complex (Figure 3.3).

The ability of extracts prepared fi'om cells at different times during development
to produce the shifted complex (Figure 3.4) correlated roughly with the timing of Q4403
expression during development (Figure 3.1). The DNA-binding activity was present by 4
h, the earliest time tested, and B-galactosidase activity from Tn5 lac Q4403 rose by 7 h
into development. DNA-binding activity reached a maximum at 12 h, whereas 8-
galactosidase activity continued to rise until 24 h, but the latter could be due to
persistence of the fiision mRN A, its continued translation, and accumulation of B-
galactosidase.

The development-specific DNA-binding activity was present in extracts prepared
from mutants defective in extracellular signaling (Figure 3 .5) despite the fact that
developmental expression of Tn5 lac Q4403 depends on signaling (Kroos and Kaiser,
1987). This could mean that the protein responsible for the DNA-binding activity is not
involved in activation of Q4403 transcription. However, it remained possible that the
protein’s ability to activate transcription was controlled not at the level of production or
DNA-binding, but rather by a modification (e.g., phosphorylation), so I continued to
characterize the DNA-binding activity.The DNA-binding activity appeared to recognize
DNA between ~41 and ~11 bp relative to the Q4403 transcriptional start site, based on
mobility shift experiments with smaller DNA fragments (Figure 3.6) and competition
experiments (data not shown). This region of the Q4403 promoter is expected to be

recognized by RNA polymerase, but it is unlikely that RNA polymerase is responsible for

133

the DNA-binding activity in the extracts because partially purified 0" RNA polymerase
(Biran and Kroos, 1997) bound to promoter-containing DNA produced a shifted complex
that migrated much more slowly through the gel (data not shown). There are many
examples of transcription factors that bind to promoters within the region recognized by
RNA polymerase (Ishihama, 1993). It remains possible that the DNA-binding activity in
the extracts is such a transcription factor. However, we were particularly interested in
identifying a protein that binds specifically to DNA between ~80 and ~72 bp because
deletion analysis had demonstrated that this region was essential for Q4403 promoter
activity in vivo. To search for such a DNA-binding activity in the extracts, the conditions
of the binding reaction were varied with respect to the monovalent (N aCl or KCl) or
divalent (MgClz) cation concentrations, the pH, the amount of nonspecific competitor
[poly(dI-dC)], and the amount of crude extract. The only shifted complex detected
migrated at the position shown in Figures 3 .2-3.6, and the conditions of the binding
reaction reported in Materials and Methods were those found to be optimal for formation
of this shifted complex.

Ifthere exists a transcription factor that binds specifically to DNA between ~80
and ~72 bp of the Q4403 promoter, there are several reasons why this protein might have
escaped detection. First, it may not be concentrated enough in the crude extract to allow
detectable binding. Proteins from crude extracts were concentrated and partially purified
using a heparin-agarose column with l M NaCl elution, followed by a DNA-cellulose
column with 0.15 M to 1 M NaCl gradient elution. In the dialyzed eluate from the
heparin-agarose column, DNA-binding activity producing a shifted complex at the

position shown in Figures 3.2-3.6 was concentrated 10-fold, but no other shifted

134

complexes were observed (data not shown). Likewise, no other shifted complexes were
observed with dialyzed DNA-cellulose column fractions (data not shown). These
experiments do not rule out the possibility that a protein binding between ~80 and ~72 bp
is present and produces a shifted complex that comigrates with the complex produced by
a protein that binds between -41 and -11 bp. Another reason why a protein binding
between ~80 and ~72 bp might have escaped detection is that this protein was susceptible
to proteolysis, despite the presence of several protease inhibitors. A third possibility is
that this protein needs another molecule in order to interact with the promoter, and this
molecule is not present or active in the crude extract.

Our goal was to understand the molecular mechanism of C-signal-dependent
activation of Q4403 transcription. As noted above, the DNA-binding activity that
recognizes DNA between -41 and -11 (Figure 3.6 and data not shown) is present in a
mutant defective in C-signaling (Figure 3.5A), which fails to express Q4403 (Kroos and
Kaiser, 1987). Nevertheless, C-signaling might modify the DNA-binding protein so that
it activates transcription, so it remained possible that this DNA-binding protein is
important for C-signal-dependent activation of gene expression. This possibility was
weakened by the finding that extracts produce similar shifted complexes with promoters
that do not depend on C-signaling for expression (Figure 3.7). Moreover, when DNA-
cellulose column fractions containing partially purified DNA-binding proteins were
tested in mobility shift assays with vegetative promoters (vegA and aphll), similar shifted
complexes were observed as with the Q4403 promoter (data not shown). These results

strongly suggest that the DNA-binding activity we have detected in developing-cell

135

extracts is not specific for C-signal-dependent promoters. Therefore, we decided not to
pursue purification and further characterization of the DNA-binding activity.

It remains possible that this DNA-binding activity represents a transcription factor
for both developmental and vegetative genes. A shifted complex was not observed with a
DNA fragment that does not contain a promoter (data not shown). Further studies would
be needed to determine whether the DNA-binding activity is indeed specific-for promoter

DNA and whether it binds to a similar region in difi‘erent promoters.

136

Summary and Perspectives

137

The multicellular developmental process of M xanthus is governed by
extracellular signaling interactions. My research focused on characterizing two
developmental genes. The aim of these studies was to investigate the regulatory
mechanisms that mediate signaling interactions and developmental gene expression.

Tn5 lac Q4514, a developmentally induced lacZ fusion originally thought to be
partially dependent on C-signaling (Kroos and Kaiser, 1987), was shown to be C-signal-
independent (chapter 2 Discussion). The regulation of the Q4514 operon was shown to be
complex and unique for M xanthus, being negatively autoregulated by one of its own
gene products, ORF1, and positively regulated by one or more putative activators that
bind to upstream DNA elements. I was unable to demonstrate binding of ORF1 protein to
the smallest Q4514 promoter DNA segment that exhibits negative autoregulation by
ORF1, and possible explanations have been discussed in chapter 2. Another possible
explanation is that the binding site for ORF1 is too close to one end of the fragment used
in the gel shift experiments, so a larger fragment could be tested.

Since multiple upstream regulatory elements are needed for fiill Q4514
expression, as demonstrated by deletion analysis, finer deletions to localize the DNA
elements that participate in regulation could be carried out. If a small DNA segment is
shown to be critical, biochemical approaches could be used to identify regulatory proteins
that interact with this DNA, as was attempted for Q4403 (chapter 3). Any proteins so
identified would be candidates for future studies aimed at comparing their expression and
activity in wild-type cells and signaling defective mutants.

Disrupting the negative autoregulation of the Q4514 operon by creating an 0111

disruption mutant caused expression from the smallest Q4514 promoter segment tested to

138

increase dramatically both during growth and development. It would be interesting to test
a promoter segment containing the upstream regulatory elements for expression in the

absence of ORF1. The results presented in chapter 2 suggest that such an experiment
might show even more dramatic effect of ORF1 on the timing and level of Q45 14
expression.

The fact that Q4514 mRNA peaked at 12-h into development whereas 8-
galactosidase activity from Tn5 lac Q4514 increased until 48-h into development is
intriguing (chapter 2 Results). No additional promoter was identified upstream by 81
nuclease protection assays, primer extension, or by an in vivo experiment in which lacZ
was integrated just upstream (i.e., at ~55) of the Q4514 promoter characterized here. One
possible explanation is that Tn5 lac Q4514 disrupts or exerts a polar efi‘ect on a gene
whose product negatively regulates Q4514 expression (chapter 2 Discussion). To address
this hypothesis, the level of Q4514 mRNA in cells containing Tn5 lac Q4514 could be
checked during development.

The DNA region between ~80 and ~72 is crucial for C-signal-dependent
expression of Q4403 (Fisseha et al., 1996). No protein binding specifically to this
sequence was identified in my experiments (chapter 3). The regulatory protein that
interacts with this region may exist at a very low level so that it is hard to observe the
protein-DNA interaction even when the extract is partially purified and concentrated by
column chromatography. Several other approaches could be tried to identify the putative
regulatory protein, including a DNA affinity column using the sequence between ~80 and

~72, using this sequence as a probe in Southwesterns, or as the target for binding in a

139

yeast one-hybrid screen, or by doing a classical genetic screen for mutants that fail to

express Tn5 lac Q4403.

140

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141

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