University of Iowa
Iowa Research Online
Theses and Dissertations
2014
A tale of two RLPAs : studies of cell division in
Escherichia coli and Pseudomonas aeruginosa
Matthew Allan Jorgenson
University of Iowa
Copyright 2014 Matthew Allan Jorgenson
This dissertation is available at Iowa Research Online: http://ir.uiowa.edu/etd/1342
Recommended Citation
Jorgenson, Matthew Allan. "A tale of two RLPAs : studies of cell division in Escherichia coli and Pseudomonas aeruginosa." PhD
(Doctor of Philosophy) thesis, University of Iowa, 2014.
http://ir.uiowa.edu/etd/1342.
Follow this and additional works at: http://ir.uiowa.edu/etd
Part of the Genetics Commons
A TALE OF TWO RLPAS: STUDIES OF CELL DIVISION IN ESCHERICHIA COLI
AND PSEUDOMONAS AERUGINOSA
by
Matthew Allan Jorgenson
A thesis submitted in partial fulfillment
of the requirements for the Doctor of
Philosophy degree in Genetics
in the Graduate College of
The University of Iowa
August 2014
Thesis Supervisor: Associate Professor David Weiss
Copyright by
MATTHEW ALLAN JORGENSON
2014
All Rights Reserved
Graduate College
The University of Iowa
Iowa City, Iowa
CERTIFICATE OF APPROVAL
_______________________
PH.D. THESIS
_______________
This is to certify that the Ph.D. thesis of
Matthew Allan Jorgenson
has been approved by the Examining Committee
for the thesis requirement for the Doctor of Philosophy
degree in Genetics at the August 2014 graduation.
Thesis Committee: ___________________________________
David Weiss, Thesis Supervisor
___________________________________
Craig Ellermeier
___________________________________
Robert Piper
___________________________________
William Moye-Rowley
___________________________________
Timothy Yahr
To my beloved Michelle.
ii
Often when you think you’re at the end of something, you’re at the beginning of
something else.
Fred Rodgers
iii
ACKNOWLEDGMENTS
It is a terrible thing to have to reduce to a few lines the importance of the
individuals in my life who have brought me to this point in my education. Know that
between these lines are found the tireless efforts, wonderful conversations, and continual
encouragements that have made all that follows possible. I would like to say to each of
you beforehand, with the deepest sincerity, that you are special and may these words
serve as a blessing for all that you have done for me.
I would like to acknowledge first David Weiss. As my mentor, I could not have
found one better. Your approach to science through critical thinking and careful planning
has played no small part in shaping the way I think. Thank you for always making
yourself available to me. Because of these, I know my time in your lab has not simply
been spent but well spent.
I want to next acknowledge my labmates. I thank Atsushi Yahashiri for being a
gracious post-doc who constantly challenged me to think differently. I also want to thank
Eric Ransom, who I could bounce an idea off of anytime and who picked me up when
things just did not want to work, as they are often prone to do in science. I count you
both as the best sort of friends to have.
I would like to thank Michael Feiss for making my first rotation such a positive
experience and introducing me to David. Because of these, I hope the Cubs win the
World Series…someday. I would also like to thank the members of my thesis committee
for lending their expertise to my projects when called upon. Your direction and advice
were invaluable and went a long way to making these stories possible.
iv
I would like to thank Pastor Tim Waldron and the members of Faith Baptist
Church in Iowa City. From my first week in Iowa, you welcomed my family into your
community and made us feel so at home when we felt so far away from our own. Your
loving kindness for our family has made all the difference. We will miss you dearly.
To my parents, I want to thank you for providing unending encouragement and
support in my life. I do not know where I would be without your model of patience, hard
work, and unwavering faith. I stand on your shoulders. And to my sister, thank you for
your constant support.
To my in-laws, you have become an integral part of my life that I could not do
without. I do not know a place where the sun shines quite so bright as in Pleasant Valley.
To John Knutson, you were always a phone call away and our conversations
about all things Minnesota sports has been an intellectual oasis.
To my precious children, Claudia and Eleanor, you make everything worth the
effort. Life was so boring before you.
Finally, to Michelle, I want you to know just how much you mean to me. There is
no other person with whom I would want to share in all the struggles of life. Thank you
for your unending love and support. Few things in life are sure; you are one of them.
Thank you, thank you all.
v
ABSTRACT
Rare lipoprotein A (RlpA) has been studied previously only in Escherichia coli,
where it localizes to the septal ring and scattered foci along the lateral wall, but null
mutants have no phenotypic change. In this thesis, we show rlpA mutants of
Pseudomonas aeruginosa form chains of short, fat cells when grown in media of low
osmotic strength. These morphological defects indicate RlpA is needed for efficient
separation of daughter cells and maintenance of rod shape. Analysis of peptidoglycan
sacculi from a ΔrlpA mutant revealed increased tetra and hexasaccharides that lack stem
peptides (hereafter called “naked glycans”). Incubation of these sacculi with purified
RlpA resulted in release of naked glycans containing 1,6-anhydro N-acetylmuramic acid
ends. RlpA did not degrade sacculi from wild-type cells unless the sacculi were
subjected to a limited digestion with an amidase to remove some of the stem peptides.
Collectively, these findings indicate RlpA is a lytic transglycosylase with a strong
preference for naked glycan strands. We propose that RlpA activity is regulated in vivo
by substrate availability, and that amidases and RlpA work in tandem to degrade
peptidoglycan in the division septum and lateral wall.
Our discovery that RlpA from P. aeruginosa is a lytic transglycosylase motivated
us to reinvestigate RlpA from E. coli. We confirmed predictions that RlpA of E. coli is
an outer membrane protein and determined its abundance to be about 600 molecules per
cell. However, multiple efforts to demonstrate that E. coli RlpA is a lytic
transglycosylase were unsuccessful and the function of this protein in E. coli remains
obscure.
vi
TABLE OF CONTENTS
LIST OF TABLES ...............................................................................................................x
LIST OF FIGURES ........................................................................................................... xi
CHAPTER
1.
INTRODUCTION ............................................................................................1
Cell division ..............................................................................................1
Peptidoglycan ............................................................................................3
Peptidoglycan synthases ...........................................................................5
Peptidoglycan hydrolases..........................................................................5
SPOR domains ..........................................................................................8
The SPOR domain proteins FtsN, DamX, and DedD .............................10
The SPOR domain protein RlpA ............................................................12
Thesis overview ......................................................................................13
2.
METHODS AND MATERIALS ...................................................................28
Media ......................................................................................................28
Strains .....................................................................................................28
Costruction of strains for P. aeruginosa studies ............................28
Construction of strains for E. coli studies ......................................29
Plasmids ..................................................................................................29
Plasmid for rescue of P. aeruginosa ΔrlpA by RlpAPa ..................29
Plasmid for rescue of P. aeruginosa ∆rlpA by RlpAEc
proteins ...........................................................................................29
Plasmids for gene knockouts in P. aeruginosa ..............................30
Plasmids for localization of RlpAPa proteins..................................30
Plasmids for localization of RlpAPa proteins with amino acid
substitutions in the DPBB domain .................................................31
Plasmid for localization of RlpAEc.................................................33
Plasmid for localization of AmiB ..................................................33
Plasmids for purification of His6-RlpA proteins ............................33
Protein purification .................................................................................34
Morphology of P. aeruginosa dacC, mltb1, rlpA,
and sltb1 mutants ....................................................................................35
Morphology of E. coli amidase and lytic transglycosylase mutants .......35
Rescue of P. aeruginosa rlpA and dacC mutants ...................................35
Scanning electron microscopy (SEM) ....................................................35
Protein localization and microscopy .......................................................36
FLIP experiments ....................................................................................36
Plasmolysis assay ....................................................................................37
Preparation of PG and labeling with RBB ..............................................37
The dye-release assay for RlpA activity .................................................38
Muropeptide analysis of PG hydrolase reactions....................................38
Renaturing gel electrophoresis for PG hydrolase activity ......................39
Western blotting ......................................................................................39
Quantification of RlpAEc protein using Western blotting .......................40
Construction of phylogenetic trees .........................................................40
vii
3.
THE BACTERIAL SEPTAL RING PROTEIN RLPA IS A LYTIC
TRANSGLYCOSYLASE THAT CONTRIBUTES TO ROD SHAPE
AND DAUGHTER CELL SEPARATION IN PSEUDOMONAS
AERUGINOSA ..............................................................................................48
Introduction .............................................................................................48
Results .....................................................................................................51
An rlpA mutant has a chaining phenotype in P. aeruginosa .........51
Septal localization of P. aeruginosa RlpA ....................................53
Low osmolarity does not induce rlpA ............................................54
PG from the ΔrlpA mutant is enriched in naked glycans ...............54
RlpA is an unusual lytic endo-transglycosylase with
specificity for glycan strands that lack stem peptides ....................56
RlpA degrades the product of amidase digestion ..........................58
Residue D157 in the DPBB is critical for lytic
transglycosylase activity ................................................................59
Evidence that RlpA is not needed for proper function of
SltB1, MltB1 or AmiB ...................................................................60
Discussion ...............................................................................................61
RlpA is a new lytic transglycosylase with an unusual
specificity for naked glycans .........................................................62
Models for how RlpA could facilitate daughter cell
separation and maitenance of rod shape ........................................63
Comparison to MltE and SpoIID ...................................................65
Potential new insights into MltA, a bacterial “expansin”, and
a protein of unknown function .......................................................65
4.
IN VIVO AND IN VITRO STUDIES SUGGEST RLPA OF
ESCHERICHIA COLI IS NOT A LYTIC TRANSGLYCOSYLASE ........106
Introduction ...........................................................................................106
Results ...................................................................................................107
RlpAEc does not rescue a P. aeruginosa ∆rlpA mutant ...............107
RlpAEc does not exhibit PG hydrolase activity in vitro ...............108
Residue D168 in the DPBB is necessary for RlpAPa function
in vivo ...........................................................................................109
A S147D substitution in the DPBB of RlpAEc is not
sufficient to restore catalytic activity ...........................................109
An rlpAEc mutation does not exhibit synthetic phenotypes
in combination with PG hydrolase mutations in E. coli ..............110
RlpAEc is an OM lipoprotein that is present at 600
molecules per cell ........................................................................111
Discussion .............................................................................................112
5.
FUTURE DIRECTIONS ..............................................................................134
E. coli ....................................................................................................134
What sequence changes are necessary to restore PG
hydrolase activity to RlpAEc?.......................................................134
What is the function of RlpAEc? ..................................................135
P. aeruginosa ........................................................................................136
What is the function of PA4485? .................................................136
viii
How do the DPBB domain and SPOR domain of RlpAPa
work together? .............................................................................136
Does RlpAPa work together with a specific amidase? .................137
What is the enzymatic specificity of RlpAPa? ..............................137
Is RlpAPa important for virulence? ..............................................138
Why salt? .....................................................................................138
REFERENCES ......................................................................................................140
ix
LIST OF TABLES
Table
1.1
The periplasmic PG hydrolases in E. coli .................................................................15
2.1
Strains used in this study ..........................................................................................42
2.2
Plasmids used in this study .......................................................................................44
2.3
Primers used in this study .........................................................................................46
3.1
Morphological parameters of a mutant lacking rlpA. ...............................................68
3.2
Functionality of various RlpA-mCherry fusion proteins ..........................................69
3.3
Muropeptide analysis of PG from Pseudomonas aeruginosa PA14 WT and
∆rlpA grown in LB0N...............................................................................................70
3.4
Amino acid and amino sugar analysis of muropeptides ...........................................71
3.5
Tandem mass spectrometry analysis of muropeptides P5, P9 and P13 ....................72
3.6
Tandem mass spectrometry analysis of muropeptides Pa and Pb ............................73
4.1
Morphological parameters of lytic transglycosylase mutants lacking rlpA in
in E. coli ..................................................................................................................115
4.2
Morphological parameters of single amidase mutants in combination with
ΔrlpA in E. coli .......................................................................................................116
4.3
Morphological parameters of double amidase mutants in combination with
ΔrlpA in E. coli .......................................................................................................117
x
LIST OF FIGURES
Figure
1.1
A partial list of the septal ring proteins of E. coli .....................................................16
1.2
Structure of the basic repeat unit of PG from E. coli ................................................18
1.3
PG structure of E. coli indicating cleavage sites for the different classes of
periplasmic PG hydrolases........................................................................................20
1.4
Lytic transglycosylase (LT) and lysozyme (LZ) activity .........................................22
1.5
The solution structure of the SPOR domain from E. coli DamX .............................24
1.6
Isolated SPOR domains localize to the septal ring and bind PG sacculi ..................26
3.1
Model of PG and RlpA function ...............................................................................74
3.2
Growth and chaining of an rlpA mutant ...................................................................76
3.3
Scanning electron microscopy of a ΔrlpA mutant of P. aeruginosa ........................78
3.4
Phenotypes associated with rlpA ..............................................................................80
3.5
Phenotypes of rlpA mutants with a SPOR domain deletion or lesions in the
DPBB domain ...........................................................................................................82
3.6
Function and localization of mutant derivatives of RlpA .........................................84
3.7
RlpA is not upregulated by low osmolarity ..............................................................86
3.8
ΔrlpA has PG alterations as compared to wild type .................................................88
3.9
RlpA is a lytic transglycosylase that cleaves naked glycan strands .........................90
3.10 RlpA digests PG sacculi from a ΔrlpA mutant .........................................................92
3.11 RlpA does not cleave isolated tetrasaccharide ..........................................................94
3.12 Amidase-treatment of PG renders it susceptible to subsequent cleavage by
His6-RlpA..................................................................................................................96
3.13 RlpA activity is potentiated by AmiD ......................................................................98
3.14 Sequence analysis of RlpA .....................................................................................100
3.15 Other PG hydrolases: SltB1, MltB1 and AmiB ......................................................102
xi
3.16 Structural comparison with MltA ...........................................................................104
4.1
Phenotypes of ΔrlpA expressing rlpAEc or mutants of rlpA with lesions in the
DPBB domain .........................................................................................................118
4.2
Function and localization of RlpA proteins ............................................................120
4.3
RlpAEc does not have PG hydrolase activity in solution ........................................122
4.4
RlpAEc does not have activity in a renaturing gel electrophoresis (zymography)
assay ........................................................................................................................124
4.5
RlpAEc is trafficked to the OM ...............................................................................126
4.6
Quantitative Western blot showing there are approximately 600 molecules of
RlpAEc per cell ........................................................................................................128
4.7
Pfam Hidden Markov model (HMM) logo of the RlpA-like DPBB domain .........130
4.8
Conservation of the putative catalytic aspartate in RlpA from different
proteobacteria..........................................................................................................132
xii
1
CHAPTER 1: INTRODUCTION
Cell division is an essential process that is required for all living beings. In the
Weiss lab, we study cell division in bacteria using primarily the Gram-negative bacterium
Escherichia coli as our model organism. In recent years, our focus has been the
relationship between cell division and metabolism of the peptidoglycan (PG) cell wall.
We embarked on this line of inquiry when we identified a new a class of cell division
proteins that contain a PG binding domain known as the SPOR domain (Arends et al.,
2010). The proteins of this class in E. coli are FtsN, DamX, DedD, and RlpA. As with
many cell division proteins, their exact biochemical function is not known. The focus of
my dissertation has been to determine the function of one of these proteins, RlpA.
Previous studies of RlpA in E. coli did not yield any promising leads until we made a
fortuitous discovery in a related Gram-negative bacterium, Pseudomonas aeruginosa.
Using a combination of genetic and biochemical analyses, we determined RlpA to be an
unusual type of PG hydrolase needed for proper daughter cell separation and cell shape in
P. aeruginosa. Curiously, however, our current evidence suggests that RlpA from E. coli
lacks PG hydrolase activity and is dispensable for daughter cell separation and cell shape.
Cell division
Cell division in E. coli involves a mother cell dividing to create two equivalent
daughter cells through the simultaneous inward growth of the three layers of the cell
envelope: the inner membrane (IM), the PG cell wall, and the outer membrane (OM).
The process of cell division requires the assembly of a septum at the midcell. Genetic
studies of cell division in E. coli date back to work done in the 1960s in which
2
researchers identified several classes of temperature sensitive mutants that were defective
in cell division but not nucleic acid synthesis (Van De Putte et al., 1964, Hirota et al.,
1968). These mutants had a normal morphology at 30°C but when shifted to 42°C grew
as long, multinucleated filaments that would eventually lyse. These mutants were given
the designation fts for filamentation temperature sensitive. Beginning in the 1990s,
investigators started applying protein localization methods to bacteria, leading to the
discovery that the Fts proteins localized to the midcell [reviewed in (Losick & Shapiro,
1999)]. In addition, new approaches for finding cell division genes revealed a host of
new proteins involved in cell division, many of them non-essential (unlike the original fts
genes, which were discovered based in part on their essentiality) [e.g., (Dai et al., 1993,
Buddelmeijer et al., 2002, Bernhardt & de Boer, 2004, Karimova et al., 2012)]. Today,
we refer to the collection of proteins that mediate cell division as the “septal ring” or
“divisome”.
There are about 30 known septal ring proteins, including 10 essential and more
than 20 non-essential proteins (Figure 1.1) [reviewed in (de Boer, 2010, Typas et al.,
2012, Lutkenhaus et al., 2012, Egan & Vollmer, 2013)]. Together, they bring about
extensive remodeling of the cell envelope that ultimately leads to daughter cell
separation. The most widely conserved and intensively studied septal ring protein is the
tubulin-like GTPase named FtsZ. The earliest recognized event in bacterial cell division
is assembly of FtsZ into a collection of short polymers at the midcell to form a structure
called the Z-ring. The Z-ring provides force to constrict the cell and serves as a landing
pad for the recruitment of other cell division proteins. The remaining septal ring proteins
serve a variety of functions: (i) Stabilization of the Z-ring and attachment to the IM
3
(FtsA, ZapA-E, and ZipA); (ii) chromosome segregation during division (FtsK); (iii)
synthesis of new PG cell wall (FtsI, FtsW, PBP1B); (iv) hydrolysis of PG to separate
daughter cells (AmiB and AmiC); (v) regulation of PG hydrolases (FtsEX, EnvC, and
NlpD); and (vi) coordinated constriction of the OM (Tol-Pal complex). However, the
process of cell division remains poorly understood as many of these septal ring proteins
are of unknown function (e.g., FtsB, FtsQ, FtsL, YmgF, FtsP, Blr, DamX, DedD, and
RlpA).
Peptidoglycan
PG (sometimes referred to as murein) forms an uninterrupted layer known as the
sacculus that surrounds most bacteria [reviewed in (Weidel & Pelzer, 1964, Turner et al.,
2014)]. In a Gram-negative bacterium like E. coli, PG is located in the periplasmic
space, which is the region between the IM and the OM. The PG layer serves two primary
functions. The first is to provide shape to the cell. Thus, purified PG sacculi retain the
shape of the organism from which they are derived. Case-in-point, a rod-shaped
bacterium like E. coli yields rod-shaped sacculi while crescent moon-shaped bacteria like
Caulobacter crescentus yield crescent moon shaped-sacculi (Vollmer et al., 2008a,
Takacs et al., 2010). The second function of PG is to stabilize the cell against the force
of turgor. Consistent with this, cells stripped of their PG layer will readily lyse unless
they are maintained in an osmotically favorable environment (e.g., high sucrose).
Remarkably, the turgor pressure experienced by E. coli remains a matter of debate, with
published estimates ranging from 30 to 500 kPa (Cayley et al., 2000, Deng et al., 2011).
This is a range of 15-fold! For comparison, the pressure of a soccer ball is ~70 kPa and a
standard car tire is ~200 kPa.
4
PG is composed of glycan strands that consist of repeating disaccharide units of
β-1,4-linked N-acetylglucosamine (NAG) and N-acetylmuramic acid (NAM) [reviewed
in (Turner et al., 2014)]. The glycans are crosslinked by short peptides that are attached
to the lactyl group of the NAM residues through an amide linkage (Figure 1.2). These
peptides contain rare D-amino acids, which protect the PG against hydrolysis by
periplasmic proteases. In E. coli, the peptides are initially synthesized as the following
pentapeptide: L-Alanine-D-γ-Glutamate-meso-diaminopimelic acid-D-Alanine-D-Alanine
(abbreviated L-Ala-D-Glu-Dpm-D-Ala-D-Ala) (Figure 1.2) (Schleifer & Kandler, 1972).
However, some amino acids are lost during assembly and maturation of the PG, so the
predominant form of the sidechain in PG isolated from E. coli is the tetrapeptide: L-AlaD-Glu-Dpm-D-Ala (Glauner,
1988). In Gram-negative bacteria, the PG sacculus is
primarily a single layer, except at sites of division, which are thought to be multilayered.
The glycan strands are arranged roughly perpendicular to the long axis of the cell
(Labischinski et al., 1991, Gan et al., 2008). Because the PG sacculus encases the cell, it
must be continuously synthesized and degraded to facilitate growth and division. This
dynamic process is orchestrated by two broad classes of enzymes known as the PG
synthases and the PG hydrolases, and their interplay must be carefully balanced to
prevent inadvertant lysis due to turgor pressure.
Biosynthesis of PG begins in the cytoplasm and leads to the formation of lipidlinked precursors composed of undecaprenyl pyrophosphoryl-NAG-NAM-pentapeptide
(also called “lipid II”) [reviewed in (van Heijenoort, 1998)]. This is the basic building
block for PG synthesis in the periplasmic space. Lipid II is transported across the IM by
flippases, whose precise identity is not completely settled. Most investigators think FtsW
5
and RodA function as lipid II flippases at the septum and lateral wall, respectively
(Mohammadi et al., 2011, Mohammadi et al., 2014, Iwaya et al., 1978, Tamaki et al.,
1980, Henriques et al., 1998). But other researchers have argued MurJ is a flippase
(Ruiz, 2008, Butler et al., 2013, Mohamed & Valvano, 2014), a proposal that has been
challenged (Fay & Dworkin, 2009, Vasudevan et al., 2009).
Peptidoglycan synthases
Following translocation into the periplasmic space, the disaccharide-pentapeptide
moiety is incorporated into PG by transglycosylation (to grow the glycan strands) and
transpeptidation (to cross-link the peptide side chains). In E. coli, the glycan strands
average 21 disaccharide units in length, and 40-60% of the peptide side chains are crosslinked (Glauner, 1988). The PG synthases responsible for these activities are anchored to
the IM and include the penicillin-binding proteins (PBPs), so named by virtue of their
affinity for penicillin and other β-lactams, an irreversible reaction that inactivates these
enzymes [reviewed in (Goffin & Ghuysen, 2002, Macheboeuf et al., 2006)]. There are
six PBPs involved in PG synthesis in E. coli (Typas et al., 2012); three of these are
bifunctional proteins with both transglycosylase and transpeptidase domains [PBP1A,
PBP1B and (possibly) PBP1C], while two are monofunctional transpeptidases (PBP2 and
PBP3). In addition, there is a monofunctional transglycosylase named MgtA.
Peptidoglycan hydrolases
The PG synthases work in concert with PG hydrolases, which are responsible for
cleaving covalent bonds in the sacculus. Unlike the PG synthases, which are few in
number, most organisms carry a large complement of PG hydrolases. For example, E.
6
coli has more than 20 periplasmic PG hydrolases (Table 1.1). Many of these enzymes
have overlapping functions, so mutations made in any one gene often elicit little or no
phenotypic change. However, elimination of several PG hydrolases will ultimately cause
a variety of morphological and division defects, which underlines the importance of these
proteins in cell shape and division (Heidrich et al., 2001, Heidrich et al., 2002,
Priyadarshini et al., 2007, Potluri et al., 2012, Singh et al., 2012).
PG hydrolases are often defined by their enzymatic specificity and, collectively,
they cleave almost every bond in PG (Figure 1.3) [reviewed in (Vollmer et al., 2008b,
van Heijenoort, 2011)]. These proteins can be grouped into three broad classes:
amidases, peptidases, and glycosidases. The soluble periplasmic N-acetylmuramyl-L-Ala
amidases, which cleave the bond between the NAM residue and the peptide side chain,
are especially important during cell division. Mutants of E. coli laking amidases grow as
chains of unseparated daughter cells (Heidrich et al., 2001, Priyadarshini et al., 2007).
Another class of PG hydrolases, the peptidases, consists of two types of enzymes, the
carboxypeptidases and endopeptidases (Table 1.1). These enzymes cleave amide bonds
between amino acids in PG and are given DD- or LD- designations based on the
stereochemistry of the bond they cleave. The carboxypeptidases cleave terminal amino
acids from the peptide side chain, thereby limiting the extent of cross-linkage in the PG.
The endopeptidases cleave between amino acids in the side chain and can therefore break
crosslinks. Some PG hydrolases possess both carboxypeptidase and endopeptidase
activity (Table 1.1). Studies of endopeptidases indicate that they are especially important
for enlargement of the PG sacculus and are referred to as the “space-makers” (Singh et
al., 2012, Hashimoto et al., 2012, Dörr et al., 2013). The final class of hydrolases
7
includes the glycosidases, enzymes that cleave the glycan backbone. There are three
types: N-acetylglucosamidases, lysozymes, and lytic transglycosylases. Of these, only
the lytic transglycosylases are found in the periplasm of E. coli. The Nacetylglucosamidases cleave the β-1,4 bond between NAG and NAM, whereas the
lysozymes and lytic transglycosylases cleave the β-1,4 bond between NAM and NAG
(Figure 1.4). It should be noted that the lytic transglycosylases are not technically true
hydrolases. Instead, they cleave between NAM and NAG in a two-step reaction that
causes formation of a 1,6-anhydroNAM product without the use of water (Holtje et al.,
1975). Mutants lacking various numbers of lytic transglycosylases often form chains of
unseparated cells and exhibit other morphological defects, indicating these proteins are
important for both cell division and cell shape (Heidrich et al., 2002, Cloud & Dillard,
2004, Monteiro et al., 2011).
In order for the cell to grow and divide, new PG must be inserted into the sacculus
by the PG synthases. To do this, the PG hydrolases are needed to make space. Indeed,
the importance of PG hydrolysis in growth and division is highlighted by the fact that E.
coli will turnover 40-50% of its PG during one generation (Goodell, 1985, Park, 1993).
As such, spatiotemporal regulation of the PG hydrolases is paramount to prevent
inadvertent lysis of the cell. One of the major questions in the field of PG metabolism is
how the various PG hydrolases are regulated.
At least four types of regulation have been described. The first involves timed
expression of PG hydrolases. For example, many bacteriophage hydrolases are thought
to be intrinsically active (Korndorfer et al., 2006, Mayer et al., 2011) and are not
produced until late in the lytic cycle, when they are needed [reviewed in (Ptashne, 2004)].
8
Similarly, in Bacillus subtilis PG hydrolases involved in sporulation are induced at the
appropriate time during spore development (Lopez-Diaz et al., 1986, Kuroda et al., 1993,
Frandsen & Stragier, 1995). However, at least for E. coli, it is not thought that PG
hydrolase production varies during the cell cycle (Arends & Weiss, 2004). The second
mechanism for regulating PG hydrolases involves activation of latent enzyme activity by
a pseudo-ABC transporter named FtsEX. FtsEX can either directly activate various PG
hydrolases or work in tandem with ancillary proteins to do the same (Sham et al., 2011,
Sham et al., 2013, Meisner et al., 2013, Yang et al., 2011, Bartual et al., 2014). The third
mechanism is related to the first and involves autoinhibition. Specifically, many bacterial
PG hydrolases appear to contain regulatory domains that occlude the active site of these
enzymes; interaction of these inhibitory domains with regulatory factors (such as FtsEX
in the septal ring) leads to conformational changes that release the enzymes from
inhibition (Yang et al., 2012, Rocaboy et al., 2013, Bartual et al., 2014). Finally, in
contrast to the protein-protein interactions described above, PG hydrolase activity has
also been shown to be regulated by substrate availability. This type of regulation is
illustrated by the interplay of SpoIID and SpoIIP, two hydrolases needed for sporulation
in B. subtilis (Lopez-Diaz et al., 1986, Frandsen & Stragier, 1995). During engulfment of
the spore, the amidase activity of SpoIIP removes stem peptides to generate naked
glycans, which in turn are digested by SpoIID, a lytic transglycosylase that can only cut
naked glycans (Morlot et al., 2010, Gutierrez et al., 2010).
SPOR domains
SPOR domains (Pfam 05036) are approximately 75 amino acids long and are
found in over 7,000 proteins in more than 2,000 species of bacteria (Pfam version 27.0)
9
(Finn et al., 2014). The SPOR domain is a PG binding domain and is named for the
founding member of this protein domain family, a cell wall amidase named CwlC that is
involved in sporulation in B. subtilis (Kuroda et al., 1993, Smith & Foster, 1995,
Mishima et al., 2005). E. coli has four SPOR domain proteins: FtsN, DamX, DedD, and
RlpA (Gerding et al., 2009, Arends et al., 2010). The structure of the SPOR domain has
been solved for CwlC, FtsN, and DamX, revealing a conserved core that consists of 4 βstrands supported on one side by two α-helices (Mishima et al., 2005, Yang et al., 2004,
Williams et al., 2013) (Figure 1.5). The SPOR domain of DamX is distinct from the
others in that it contains an additional alpha helix at the C-terminus that associates with
the opposite side of the β-sheet and is important for function (Williams et al., 2013)
(Figure 1.5). Mutagenesis studies indicate that the residues important for PG binding are
found in the β-sheet (Williams et al., 2013, Duncan et al., 2013).
Though initially described to be involved in sporulation, most SPOR domain
proteins are now thought to be involved in cell division, highlighting the interplay
between cell division and PG metabolism (Möll & Thanbichler, 2009, Gerding et al.,
2009, Arends et al., 2010). Isolated SPOR domains localize to the septal ring in vivo and
bind PG in vitro (Gerding et al., 2009, Arends et al., 2010). Thus, residues important for
septal ring localization are also important for PG binding (Williams et al., 2013, Duncan
et al., 2013). This observation argues SPOR domains localize by binding to some form
of PG that is enriched in division septa. This mechanism of localization is unusual in that
most septal ring proteins are known to localize to the midcell through a cascade of
protein-protein interactions [reviewed in (Buddelmeijer & Beckwith, 2002)]. Further
support for the notion that SPOR domains localize by binding to septal PG rather than
10
septal proteins comes from the observation that SPOR domains from AQ1897 (Aquifex
aeolicus) and CHU2221 (Cytophaga hutchinsonii) localize to the septal ring when
expressed in E. coli, despite those organisms being very distant relatives of E. coli
(Arends et al., 2010).
The chemical features of septal PG recognized by SPOR domains are not well
understood, but two lines of evidence suggest binding is to naked glycan strands. First,
the SPOR domain of FtsN was shown to bind naked glycans that were longer than 25
disaccharide units (Ursinus et al., 2004). Second, the SPOR domain of FtsN does not
localize to sites of division in an E. coli mutant lacking three cell wall amidases
(∆amiABC), two of which specifically localize to the septal ring (AmiB and AmiC)
(Gerding et al., 2009). However, naked glycans have not been observed in wild type
cells, so their importance in vivo remains uncertain (Glauner, 1988, de Jonge et al., 1989,
Evans et al., 2013). Naked glycans have been reported, though, when cells are
genetically manipulated (Gilmore et al., 2004).
The SPOR domain proteins FtsN, DamX, and DedD
Like many septal ring proteins, the precise function of the SPOR domain proteins
is essentially unknown. A major challenge for the future will be to determine the
biochemical function of these proteins and how they work together to facilitate cell
division. FtsN, DamX, and DedD are alike in that they have relatively similar protein
architectures (Figure 1.1). Each is a bitopic IM protein with a cytoplasmic domain, a
single transmembrane helix, and a large periplasmic domain, the last 75 amino acids of
which comprises the SPOR domain.
11
FtsN is an essential protein (Dai et al., 1993) and is the most studied of the SPOR
domain proteins. ftsN is a multicopy suppressor of several cell division mutants, which
has led to the suggestion that it functions to stabilize the septal ring (Dai et al., 1993,
Draper et al., 1998, Geissler & Margolin, 2005, Reddy, 2007). Studies of FtsN indicate
that it localizes concurrent with the onset of constriction, suggesting FtsN might trigger
cytokinesis (Gerding et al., 2009). There seem to be conflicting ideas for how FtsN
might provoke constriction. On the one hand, the critical region of FtsN is in the
periplasmic domain [but does not include the SPOR domain (see below)], suggesting
FtsN might allosterically activate septal PG synthesis by PBP1B or FtsI (Müller et al.,
2007, Gerding et al., 2009). On the other hand, the cytoplasmic domain of FtsN was
recently shown to interact with FtsA (Busiek et al., 2012, Busiek & Margolin, 2014).
Because FtsA modulates assembly of FtsZ, this finding suggests FtsN might trigger
constriction of the Z-ring. Interestingly, the SPOR domain of FtsN is not required for its
function during cell division, although deletion derivatives that lack the SPOR domain do
not localize very efficiently (Ursinus et al., 2004, Möll & Thanbichler, 2009, Gerding et
al., 2009). Apparently small amounts of septal FtsN are sufficient for cytokinesis.
Less is known about the non-essential SPOR domain proteins. Mutants of damX
do not have division defects but are more sensitive to bile salts (Gerding et al., 2009,
Arends et al., 2010, Lopez-Garrido & Casadesus, 2010). Overproduction of DamX
inhibits cell division (Lyngstadaas et al., 1995), and DamX antagonizes the function of
the essential septal ring protein FtsQ (Arends et al., 2010). Cells containing a dedD null
mutation are slightly elongated and have a mild chaining defect (Gerding et al., 2009,
Arends et al., 2010). In addition, dedD has synthetic phenotypes with both ftsN and
12
damX, which suggests these proteins have overlapping functions (Gerding et al., 2009,
Arends et al., 2010).
The SPOR domain protein RlpA
The fourth E. coli SPOR domain protein is called rare lipoprotein A (RlpA) and
may be the most enigmatic given its pedigree. RlpA is 362 amino acids long, has a type
II signal sequence, and is predicted to be trafficked to the OM by virtue of its lipobox
motif (unlike FtsN, DamX, and DedD, which are located in the IM) (Takase et al., 1987,
Seydel et al., 1999). RlpA has a C-terminal SPOR domain, and localizes strongly to the
septal ring, although small amounts of the protein also localize to scattered foci along the
lateral wall (Gerding et al., 2009, Arends et al., 2010). This localization pattern suggests
RlpA is involved in both division and elongation. In addition to the SPOR domain, RlpA
contains an “RlpA-like double-psi β barrel domain” (Pfam 03330). The DPBB domain is
annotated in the Pfam database as usually being enzymatic, though it has different
activities in different proteins (Castillo et al., 1999). For example, the E. coli protein
MltA is a lytic transglycosylase, while another E. coli protein, PanD, is an aspartate 1decarboxylase required to synthesize β–alanine from L-aspartate (Ursinus & Höltje,
1994, Cronan, 1980). The DPBB domain of RlpA is detected not by a BLAST search but
by using protein modeling programs such as the Protein Homology/analogY Recognition
Engine (PHYRE) (Kelley & Sternberg, 2009). When threaded, the DPBB of RlpA
showed the greatest similarity to a protein of unknown function in P. aeruginosa
(PA4485). The DPBB of RlpA showed weaker structural similarity to MltA from E. coli
and to several cellulose-binding proteins called expansins, which are found in plants and
in bacteria (Sampedro & Cosgrove, 2005, Kerff et al., 2008). Expansins are not thought
13
to be enzymatic. The distant homology of RlpA to MltA and expansins suggests a
connection to carbohydrates, but whether the protein has enzymatic activity is not clear.
RlpA is widely conserved, with over 5000 examples in the Pfam database (Finn et
al., 2014). However, only 1400 of these hypothetical proteins have a SPOR domain. The
conservation of RlpA argues it plays an important role in bacterial physiology. However,
rlpA mutants of E. coli do not have division or growth phenotypes, even in combination
with deletions of damX and dedD (Gerding et al., 2009, Arends et al., 2010). Two
studies reported rlpA mutants have phenotypes that might reflect altered membrane
permeability (Nichols et al., 2011, Paradis-Bleau et al., 2014); however, these
phenotypes are subtle and difficult to interpret with respect to the function of the protein.
Because of this, studies of RlpA have been at an impasse.
Thesis overview
Chapter 2 contains a detailed description of the methods used during the course of
my investigations. Chapter 3 describes the efforts to uncover the function of RlpA in P.
aeruginosa. Briefly, having failed to turn up a tell-tale phenotype for rlpA mutants in E.
coli, we decided to study rlpA in P. aeruginosa because a mariner transposon library had
been previously constructed in the PA14 background (Liberati et al., 2006). We
discovered that an rlpA::Tn mutant had a division defect when grown in media of low
osmotic strength, forming chains of short, fat cells. Using a series of genetic and
biochemical analyses, we went on to show that RlpA is an unusual type of lytic
transglycosylase whose activity is potentiated by cell wall amidases. These findings are
important in that they (i) assign a function to RlpA, (ii) illustrate one of the ways in
which PG hydrolases are regulated in the cell (i.e. substrate availability), and (iii)
14
demonstrate how PG hydrolases work together to effect PG turnover in the cell
(sequential cutting). Chapter 4 returns to studies of RlpA in E. coli. In contrast to the
situation in P. aeruginosa, our findings argue that E. coli RlpA probably does not have
PG hydrolase activity. Finally, chapter 5 describes future directions for continued studies
of RlpA.
15
Table 1.1.The periplasmic PG hydrolases in E. colia
Protein
Gene
Localizationb
NAM-L-alanine
amidases
AmiA
AmiB
AmiC
AmiD
amiA
amiB
amiC
amiD
P
P
P
OM
1,6-anhydro-NAM-Lalanine amidase
AmiD
amiD
OM
dacB
yfeW
dacA
dacC
dacD
mepS
mepA
mepH
mepM
mepS
dacB
pbpG
mepA
pbpG
slt
mltA
mltB
mltC
mltD
mltE
mltF
IM
IM and P
IM
IM
IM
OM
P
IM/P?
IM?
OM
IM
P
P
P
P
OM
OM
OM
OM
OM
OM
Enzyme
PBP4
PBP4B
DD-Carboxypeptidases
PBP5
PBP6
PBP6B
LD-Carboxypeptidase
MepS
MepA
MepH
MepM
DD-Endopeptidases
MepS
PBP4
PBP7
MepA
LD-Endopeptidases
PBP7
Slt70
MltA
MltB
Lytic
MltC
Transglycosylases
MltD
MltE
MltF
a
Table adapted from (Typas et al., 2012).
b
IM, inner membrane; P, periplasm; OM, outer membrane.
16
Figure 1.1. A partial list of the septal ring proteins of E. coli. Filamentation temperature
sensitive (Fts) proteins are given by single letter designations. OM, outer membrane; PG,
peptidoglycan cell wall; IM, inner membrane. Figure modified from (Goehring &
Beckwith, 2005).
17
18
Figure 1.2. Structure of the basic repeat unit of PG from E. coli. The glycan strands
consists of alternating β-1,4 linked N-acetylglucosamine (NAG) and N-acetylmuramic
acid (NAM) residues. The glycosidic bond is drawn in blue. Shown is a PG monomer
with a basic pentapeptide side chain. Amino acids are connected through amide bonds
drawn in red. Ala, alanine; Glu, glutamate; Dpm, meso-diaminopimelic acid. D- and Lrefer to optical activity of the amino acid isomer. Modified from (Vollmer et al., 2008a).
19
NAG
NAM
L-Ala
D-Glu
Dpm
D-Ala
D-Ala
20
Figure 1.3. PG structure of E. coli indicating cleavage sites for the different classes of
periplasmic PG hydrolases. Ami, N-acetylmuramyl-L-alanine amidase; aAmi, 1,6anhydro N-acetylmuramyl-L-alanine amidase; CP, carboxypeptidase; EP, endopeptidase.
DD-, LD- and DL- refer to the stereochemistry of the bond. Figure modified from
(Vollmer et al., 2008b).
21
DD-CP
LT
Dpm
|
D-Glu
|
L-Ala
|
D-Ala
|
D-Ala
|
Dpm
|
D-Glu
|
L-Ala
|
aAmi
– – – NAG – NAM – NAG – NAM – NAG – NAM – NAG – aNAM
|
|
Ami
LD-CP
L-Ala
|
D-Glu DD-EP
|
Dpm D-Ala
|
|
D-Ala Dpm
|
D-Glu
|
L-Ala
|
LT
L-Ala
|
D-Glu
LD-EP
|
Dpm
|
D-Ala Dpm
|
D-Glu
|
L-Ala
|
D-Ala
|
D-Ala
|
Dpm
|
D-Glu
|
L-Ala
|
– – – NAG – NAM – NAG – NAM – NAG – NAM –NAG – NAM – NAG – NAM – – –
|
|
Ami
L-Ala
|
D-Glu
|
Dpm
DL-EP
L-Ala
|
D-Glu
|
Dpm
|
D-Ala
22
Figure 1.4. Lytic transglycosylase (LT) and lysozyme (LZ) activity. Cleavage of the
glycosidic bond between NAM and NAG by LTs (top) or LZs (bottom). The former
results in the formation of a nonreducing 1,6-anhdyro NAM residue at the end. Figure
modified from (Vollmer et al., 2008b).
23
NAG
1,6-aNAM
NAG
1,6-aNAM
NAG
1,6-aNAM
NAG
1,6-aNAM
LT
NAG
NAM
LZ
NAG
NAM
24
Figure 1.5. The solution structure of the SPOR domain from E. coli DamX. α-helices and
β-sheets are numbered from the N-terminus to the C-terminus. Figure was modified from
PDB: 2LFV (Williams et al., 2013) in Pymol (Delano, 2002).
25
α2b
α1
α2a
β1
β4
β3
β2
N
α3
C
26
Figure 1.6. Isolated SPOR domains localize to the septal ring and bind PG sacculi. (A)
Fluorescence micrographs of E. coli cells expressing the indicated GFP fusions to
isolated SPOR domains. Bar = 5 µm. (B) PG binding assay. Purified E. coli sacculi
were incubated with purified proteins and pelleted by ultracentrifugation. Pellets were
then washed and ultracentrifuged again. Samples of the supernatant, wash, and pellet
fractions were analyzed by SDS-PAGE and Coomassie staining to determine what
fraction of the total protein remained in the pellet (bound PG). Bars represent the mean
and standard deviation of three independent experiments. FtsZ and MBP (maltose
binding protein) were used as negative controls. Modified from (Arends et al., 2010).
27
A
B
28
CHAPTER 2: MATERIALS AND METHODS
Note that RlpA from P. aeruginosa and E. coli will be denoted as RlpAPa or
RlpAEc, respectively, when necessary.
Media
Unless otherwise noted, E. coli and P. aeruginosa strains were grown in LuriaBertani (LB) media containing 0.5% yeast extract, 1% tryptone, and 1% NaCl. LB
lacking NaCl is referred to as LB0N. Plates contained 1.5% agar. When necessary,
ampicillin (Amp), carbenicillin (Carb), gentamicin (Gent), irgasan (Irg), and kanamycin
(Kan) were used at 200, 300, 100, 25 and 40 µg/mL, respectively.
Strains
All strains used in this study are listed in Table 2.1. All P. aeruginosa strains
used for in vivo experiments are derivatives of UCBPP- PA14. All E. coli strains used
for in vivo experiments are derivatives of MG1655 (Guyer et al., 1981).
Construction of strains for P. aeruginosa studies. pEXG2 derivatives were
transferred from derivatives of E. coli strain SM10 to wild type PA14 by conjugation as
described (Schweizer, 1992) except that Irg was used to counter select against the E. coli
donor strain because a ΔrlpA mutant is not viable on the (low osmolarity) minimal-citrate
media often used to counter select E. coli after such matings. Resolution of the cointegrant was selected for on LB0N plates containing 5% sucrose (~150 mM, which
allows for growth of the ΔrlpA mutant in the absence of NaCl). Gene knockouts were
made in the wild type (MJ1) background. Gene knock-ins were made in the ΔrlpA
(MJ24) background.
29
Construction of strains for E. coli studies. Deletion alleles were obtained from
Keio collection (Baba et al., 2006) strains JW2428 (amiA::kan), JW5449 (amiC::kan),
JW2784 (mltA::kan), JW5481 (mltC::kan), JW5018 (mltD::kan), JW5821 (mltE::kan),
and JW4355 (slt::kan). Eviction of antibiotic markers by pCP20 and P1-mediated
transduction were done as previously described (Datsenko & Wanner, 2000, Miller,
1972). Selection for pCP20 was at 50 µg/mL Amp for lytic transglycosylase deletion
alleles because these strains were unusually Amp sensitive.
Plasmids
All plasmids and primers used in this study are listed in Tables 2.2 and 2.3. All
plasmids used for P. aeruginosa in vivo experiments are derivatives of pJN105 (Newman
& Fuqua, 1999).
Plasmid for rescue of P. aeruginosa ΔrlpA by RlpAPa. pDSW1398 (PBAD::rlpA)
was constructed by amplifying rlpA from PA14 chromosomal DNA with primers P1603
and P1604. The 1205 bp product was digested with EcoRI and XbaI and ligated to the
same sites of pJN105 to create the desired PBAD::rlpA construct. Expression of rlpA from
pDSW1398 did not require arabinose induction.
Plasmids for rescue of P. aeruginosa ∆rlpA by RlpAEc proteins. A multistep
PCR procedure involving megapriming was used to generate a fusion between the type II
signal sequence of rlpAPa and codons 18-362 of rlpAEc, which code for the mature
(secreted) protein of RlpAEc. Primers P1770 and P1773 were used to amplify upstream
sequence and codons 1-26 of rlpAPa from pDSW1518. P1773 has 22 bp of homology to
rlpAEc beginning at the sequence coding for the cysteine at residue 18. The 270 bp
product was isolated by PCR column purification (Qiagen) and used in a subsequent
30
reaction with P1774 to amplify the rest of rlpAEc (codons 18-362) from pDSW930. The
1301 bp product was cut with EcoRI and XbaI and ligated to the same sites of
pDSW1518 to produce pDSW1554 (pJN105::rlpAEc-mCherry).
RlpAEc with a S147D substitution was synthesized as a gBlock gene fragment
(Integrated DNA Technologies). Primers P1967 and P1968 were then used to amplify
rlpAEc(S147D). The 827 bp product was cut with EcoRI and XmnI and ligated to the
same sites of pDSW1554 to make pDSW1695 [pJN105::rlpAEc(S147D) –mCherry].
Plasmids for gene knockouts in P. aeruginosa. In-frame deletions were
constructed essentially as previously described using the pEXG2 vector (Schweizer,
1992, Rietsch et al., 2005). pDSW1385 (pEXG2::‘sltb1 ΔrlpA dacC’) was constructed
by amplifying ~1 Kb of upstream sequence plus the first 8 codons of rlpA with primers
P1507 and P1474. Similarly, the last 8 codons and ~1 Kb of sequence downstream of
rlpA were amplified with primers P1475 and P1508. The 975 and 994 bp products,
respectively, were cut with XbaI and ligated to each other to make a 1955 bp product,
which was further amplified using primers P1507 and P1508. The 1955 bp product was
cut with HindIII and MfeI and ligated to pEXG2 cut with HindIII and EcoRI. Similar
procedures were used to construct pDSW1490 (pEXG2::‘rodA Δsltb1 rlpA’) and
pDSW1516 (pEXG2::‘PA14_57740 Δmltb1 cysD’) using the following primers: P1702P1705 (pDSW1490) and P1713-1716 (pDSW1516).
Plasmids for localization of RlpAPa proteins. To construct an RlpAPa-mCherry
fusion, primers P1599 and P1600 were used to amplify rlpA from PA14 chromosomal
DNA. The 1047 bp product was cut with EcoRI and XbaI and ligated to the same sites of
pDSW913 (P206::MCS-mCherry) to create pDSW1399. Similarly, to construct a SPOR
31
deletion mutant of RlpA for localization studies, primers P1599 and P1708 were used to
amplify rlpA(Δ269-341) from PA14 chromosomal DNA. The 828 bp product was cut
with XbaI and EcoRI and ligated to the same site of pDSW913 to produce pDSW1497.
For localization studies in P. aeruginosa, rlpA-mCherry was recombined onto the
chromosome using procedures similar to those used to make gene deletions. pDSW1489
(pEXG2::‘rlpA-mCherry dacC’) was constructed by amplifying rlpA-mCherry from
pDSW1399 using primers P1680 and P1681. In a subsequent reaction, ~1 Kb of
sequence downstream of rlpA was amplified with P1682 and P1683. The 1761 bp and
1018 bp products, respectively, were cut with MfeI, ligated to each, and further amplified
using primers P1680 and P1683. The 2.8 Kb product was digested with HindIII and
KpnI and ligated to the same sites of pEXG2 to make pDSW1489. A similar procedure
was used to generate a pEXG2 derivative for recombining the rlpA SPOR deletion fusion
[rlpA(Δ269-341)-mCherry] onto the chromosome of P. aeruginosa. pDSW1504
[pEXG2::‘rlpA(Δ269-341)-mCherry dacC’] was constructed by amplifying rlpA(Δ269341)-mCherry from pDSW1497 using primers P1680 and P1681. The 1542 bp product
was cut with HindIII and MfeI and ligated to the same sites of pDSW1489 to produce
pDSW1504.
Plasmids for localization of RlpAPa proteins with amino acid substitutions in
the DPBB domain. Amino acid substitutions in the DPBB domain of RlpA were
generated using a multistep PCR procedure involving megapriming (Kwok et al., 1994).
To introduce substitutions, rlpA-mCherry was amplified from pDSW1399 with primers
P1599 and P1727. The 1764 bp product was cut with AatII and SacI, then ligated to the
same sites of pDSW1398 to produce the vector pDSW1518. Amino acid substitutions in
32
the DPBB domain of rlpA were then introduced by megapriming. For example, a D157N
substitution in the DPBB of rlpA was constructed by amplifying rlpA from pDSW1518
with primers P1754 and P1781. P1781 has a sequence change at the codon for D157.
The 315 bp product was isolated by PCR column purification and used in a subsequent
reaction with primer P1755 to produce full length rlpA (with the D157N substitution)
from pDSW1518. The 870 bp product was cut with AatII and XbaI and ligated to the
same sites of pDSW1518 to produce pDSW1545. Similar procedures were used to
introduce substitutions at other residues using the following primers in place of P1781:
P1756 (E120A) to make pDSW1519, P1758 (D123A) to make pDSW1520, P1760
(H131A) to make pDSW1537, P1957 (D168N) to make pDSW1676, and P1959 (D168S)
to make pDSW1694.
For functional studies in P. aeruginosa, pEXG2 derivatives containing rlpA
variants with amino acid substitutions in the DPBB domain were generated for
recombination onto the chromosome of P. aeruginosa. pDSW1614 (pEXG2::‘sltb1 rlpAmCherry dacC’) was constructed by amplifying ~1 Kb of sequence upstream of rlpA
from PA14 chromosomal DNA using primers P1821 and P1822. The 1208 bp product
was cut with BamHI and HindIII, and ligated to the same sites of pDSW1489 to make
pDSW1614. pDSW1614 was then used as a destination vector for mutants of rlpA with
substitutions in the DPBB domain. pDSW1615 [pEXG2::‘sltb1 rlpA(E120A)-mCherry
dacC’] was constructed by amplifying rlpA with an E120A substitution from pDSW1519
using primers P1823 and P1824. The 556 bp product was cut with BamHI and NotI and
ligated to the same sites of pDSW1614. Similar procedures were used to introduce other
rlpA variants with amino acid substitutions in the DPBB domain using the following
33
plasmids as template: pDSW1520 (D123A) to make pDSW1616, pDSW1537 (H131A) to
make pDSW1617 and pDSW1545 (D157N) to make pDSW1619.
Plasmid for localization of RlpAEc. RlpAEc-mCherry was previously
constructed by David Weiss. Briefly, rlpA was amplified from E. coli MG1655
chromosomal DNA using primers P1140 and P1141. The 1113 bp product was cut with
EcoRI and XbaI and ligated to the same sites of pDSW912 to make pDSW930.
Plasmid for localization of AmiB. To construct an AmiB-mCherry fusion,
primers P1805 and P1806 were used to amplify amiB from PA14 chromosomal DNA.
The 1483 bp product was cut with EcoRI and XbaI and ligated to the same sites of
pDSW1518 to produce pDSW1635.
Plasmids for purification of His6-RlpA proteins. Plasmids for overproducing
hexahistidine (His6-) tagged RlpA variants are derivatives of pQE-80L (Qiagen). To
overproduce RlpAPa with an N-terminal His-tag, rlpAPa was amplified from PA14
chromosomal DNA with primers P1787 and P1711. The 962 bp product was cut with
BclI and HindIII and ligated to pQE-80L cut with BamHI and HindIII to make
pDSW1557. Similar procedures were used to clone rlpA variants with amino acid
substitutions [pDSW1600 (D157N), pDSW1601 (E120A), pDSW1604 (D123A) and
pDSW1606 (H131A) using plasmids pDSW1545, pDSW1519, pDSW1520 and
pDSW1537, respectively, as template]. Purification constructs contain amino acids 28341 of rlpA and the sequence MRGSHHHHHHGS at the N-terminus.
To overproduce RlpAEc with an N-terminal His-tag, rlpA was amplified from E.
coli chromosomal DNA using primers P1375 and P1128. The 1043 bp product was cut
with BglII and HindIII and ligated to pQE-80L cut with BamHI and HindIII to make
34
pDSW1132. To overproduce RlpAEc with a S147D substitution, rlpAEc was amplified
with P1375 and P1951 from pDSW1132. P1951 has a sequence change at the codon for
S147. The 408 bp product was used as a megaprimer in a subsequent reaction with
P1128. The 1043 bp product was cut with BglII and HindIII and ligated to pQE-80L cut
with BamHI and HindIII to make pDSW1674. The sequence for the affinity tag in both
cases was MRGSHHHHHHGSNNN.
Protein purification
Wild type and mutant His6-RlpA proteins were overproduced in E. coli BL21 and
purified at 4°C by cobalt affinity chromatography per the manufacturer’s instructions
(Clontech). Cells were grown at 30°C to an OD600 ~0.5 and protein production was
induced with 1 mM isopropyl-ß-D-thiogalactoside (IPTG) for three hours. The purified
proteins were dialyzed into storage buffer (25 mM HEPES, 150 mM NaCl, 5% glycerol,
pH 7.0) at 4°C and aliquots were stored at -80°C until needed. Typical yields were 10
mg from a 500 mL culture as determined by UV-VIS spectrometry using a NanoDrop1000 spectrophotometer (Thermo Scientific) and purity was judged to be ~95% by
sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). The extinction
coefficient used to determine the amount of His6-RlpAPa and His6-RlpAEc by nanodrop
was 46,870 and 16,055, respectively. His6-AmiD was purified essentially as described
and stored at -80°C (Uehara & Park, 2007). The purity was ~95% as judged by SDSPAGE.
35
Morphology of P. aeruginosa dacC, mltb1, rlpA, and sltb1 mutants
Overnight cultures grown in LB were adjusted to an OD600 ~0.05 (~1:100
dilution) in the same medium and grown to an OD600 ~1.0 at 37°C. Cultures were then
diluted to an OD600 ~0.05 in LB0N medium and grown to an OD600 ~0.5 at which point
the cells were fixed as described except that glutaraldehyde was omitted (Pogliano et al.,
1995). Cells were stained with the membrane dye FM4-64 (Invitrogen) to better
visualize chaining.
Morphology of E. coli amidase and lytic transglycosylase mutants
Overnight cultures grown in LB were diluted 1:2000 in the same medium and
grown to an OD600 ~0.5-0.6 at 37°C. Cells were then fixed and stained with the
membrane dye FM4-64 as above. Morphological parameters from double mutants of
∆amiA ∆amiC were determined from overnight cultures grown in LB at 37°C.
Rescue of P. aeruginosa rlpA and dacC mutants
Overnight cultures were adjusted to OD600 = 1.0 and plating efficiency was
assessed by spotting tenfold serial dilutions onto LB or LB0N plates (Gent was added to
plates for strains containing plasmids) (Arends et al., 2010). Plates were incubated for 18
hours at 37°C and then photographed.
Scanning electron microscopy (SEM)
Overnight cultures grown in LB were matched to an OD600 ~0.01 in the same
medium and grown to an OD600 ~1.0 at 37°C. Cultures were then diluted to an OD600
~0.1 in LB0N medium and grown to an OD600 ~0.7 at which point the cells were fixed
36
with 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer and prepared essentially as
previously described (Hsiao et al., 2011). Samples were examined by an S-4800 field
emission scanning electron microscope (Hitachi High Technologies America Inc.). All
electron microscopy was performed at the University of Iowa Central Microscopy
Research Facility.
Protein localization and microscopy
Strains expressing mCherry fusion proteins were grown overnight in LB, adjusted
to an OD600 = 0.02 (~1:200 dilution) in the same medium and grown at 37°C to an OD600
~1.0. Cultures were then diluted to an OD600 ~0.1 in LB0N and grown to an OD600 ~0.5
after which 4 μl were spotted onto 1% agarose pads and visualized by phase-contrast and
fluorescence microscopy (Tarry et al., 2009). For imaging mCherry fusions we used a
filter for Texas Red (Chroma no. U-N41004). Our microscope, camera and software
have been previously described (Mercer & Weiss, 2002).
FLIP experiments
Cells expressing cytoplasmic GFP from pMRP9-1 (Davies et al., 1998) were
grown overnight at 37°C in LB containing carbenicillin and diluted 1:100 in the same
medium and grown to an OD600 ~1.0. Cultures were then diluted 1:10 in LB0N
containing carbenicillin and grown to an OD600 = 0.5 after which 4 µL were spotted onto
1% agarose pads. A coverslip was placed over the sample and sealed with nail polish.
Fluorescence loss in photobleaching (FLIP) was then performed using a Zeiss LSM 510
confocal microscope essentially as described (Priyadarshini et al., 2007). All confocal
37
microscopy was performed at the University of Iowa Central Microscopy Research
Facility.
Plasmolysis assay
Plasmolysis was done essentially as previously described (Lewenza et al., 2006).
Cells grown in LB Amp overnight were diluted 1:100 in the same medium containing
100 µM IPTG to induce expression of fusion proteins. Cells were grown to an OD600
~0.5 after which 1 mL of culture was harvested and resuspended in 1 mL of LB or 1 mL
of plasmolysis solution (15% sucrose, 25 mM HEPES [pH 7.4]). Cells were pelleted
again and resuspended in the same medium. 4 µL were then spotted onto 1% agarose
pads or 1% agarose with 15% sucrose (to maintain plasmolysis) and visualized by phasecontrast and fluorescence microscopy.
Preparation of PG and labeling with RBB
Whole PG sacculi were isolated from 1-liter cultures as previously described
(Arends et al., 2010). Overnight cultures of P. aeruginosa strains grown in LB were
diluted to an OD600 ~0.05 in the same medium and grown to an OD600 ~1.0 at 37°C.
Cultures were then diluted to an OD600 ~0.05 in LB0N and grown for three hours (OD600
~0.5-0.6) before harvest. Remazol Brilliant Blue (RBB) labeling of PG was performed
essentially as previously described (Uehara et al., 2010, Zhou et al., 1988). Typically,
purified sacculi from 1 liter of culture were incubated in a volume of 1 mL of 20 mM
RBB in 0.25 M NaOH overnight at 37°C. Reactions were neutralized by the addition of
HCl and RBB-PG was collected by centrifugation at 18,000 x g for 15 min. Pellets were
38
washed repeatedly with water until the supernatants were colorless. RBB-labeled sacculi
were stored in water at 4°C until needed.
The dye-release assay for RlpA activity
A standard 100 µL reaction mixture contained PBS buffer (137 mM NaCl, 3 mM
KCl, 9 mM NaH2PO4 and 2 mM KH2PO4, pH 7.4), 10 µL of RBB-labeled PG, and 4 µM
lysozyme or His6-RlpA (as indicated). Reactions were incubated for 18 hours at 30°C or
37°C, then stopped by centrifugation at 18,000 x g for 10 minutes. (We did not boil
reaction mixtures because we found that caused a little bit of dye-release and thus
interfered with measuring low activities.) The supernatants were removed and their
absorbance was measured at 595 nm using a Beckman Coulter DU60 Spectrophotometer.
To test whether limited digestion with an amidase potentiated subsequent digestion by
His6-RlpA, we first subjected dye-labeled sacculi from wild-type E. coli to a limited
digestion with His6-AmiD. Reaction mixtures (1 mL) contained PBS buffer, 300 μL of
RBB-PG, and 1 μM His6-AmiD. Digestion was allowed to proceed for 18 hours at 37°C,
and then terminated by heating to 95°C for 10 min. Sacculi were recovered by
centrifugation and washed repeatedly with water until the supernatants were colorless (~4
washes). His6-AmiD-treated RBB-PG was then suspended in 300 μL water, and used in
assays as described above. As controls, sacculi were digested overnight with 1 μM His6RlpA or protein was omitted.
Muropeptide analysis of PG hydrolase reactions
Sacculi were incubated with wild type or mutant His6-RlpA protein (4 µM) for 2
hours at 37°C in PBS. Reactions were terminated by heating to 95°C for 5 minutes.
39
Reaction mixtures were centrifuged at 18,000 x g for 15 minutes and the supernatant was
separated from the pellet. The PG in both fractions was then prepared for highperformance liquid chromatography (HPLC) as described (Popham et al., 1996a).
Purified muropeptides were identified by amino acid/amino sugar analyses and mass
spectrometry as previously described (Popham et al., 1996b).
Renaturing gel electrophoresis for PG hydrolase activity
Zymography was performed essentially as previously described (Gutierrez et al.,
2010). Purified proteins were subjected to 10% SDS-PAGE gels with or without 0.5%
Micrococcus lysodeikticus cells (Sigma). Gels were briefly washed in water and then
gently shaken in 300 mL of renaturing solution (25 mM Tris-HCl, 1% Triton X-100 [pH
7.2]) at 37°C overnight. Gels were imaged against a black background and then stained
with 0.01% methylene blue in 0.01% KOH for 3 hours. Gels were rinsed extensively
with water and imaged again.
Western blotting
Western blotting was performed as previously described (Arends et al., 2010).
Briefly, cells from 1 mL of culture grown to an OD600 ~0.5 were centrifuged,
resuspended in 200 µL of Laemmli sample buffer containing 5% β- mercaptoethanol and
boiled for 10 minutes. 10 µL aliquots were subjected to SDS-PAGE (10%
polyacrylamide). Proteins were transferred onto nitrocellulose membranes and detected
using standard methods. Rabbit anti-RFP serum (a gift from L. Shapiro) was used at a
1:10,000 dilution. The secondary antibody (used at a 1:8,000 dilution) was horseradish
peroxidase-conjugated goat anti-rabbit antibody (Thermo Scientific) and detection was
40
with SuperSignal West Pico Chemiluminescent Substrate (Thermo Scientific). Blots
were visualized using a Fujifilm LAS-1000 imager.
Quantification of RlpAEc protein using Western blotting
Quantitative Western blotting was performed as previously described (Weiss et
al., 1997). Overnight cultures of EC251 grown in LB were diluted 1:200 in the same
medium and grown at 37°C to an OD600 ~0.5 before harvest. The number of cells was
determined by serial dilutions and plating on LB. Western blotting was used to compare
the amount of RlpAEc in the cell to a standard curve of purified His6-RlpAEc added to an
E. coli ∆rlpA extract (EC3183). The amount of His6-RlpAEc was determined by
absorbance at 280 nm using a predicted molecular mass of 35.7 kDa and an extinction
coefficient of 16,055 as determined by the ExPASy ProtParam website
[http://web.expasy.org/protparam/]. Blots were visualized as above and the Quant
software was used to quantify fluorescent signals.
Construction of phylogenetic trees
Phylogenetic trees were adapted from previous reports (Emerson et al., 2007,
Baumler et al., 2013). Conservation of the putative catalytic residue of RlpA from
different proteobacteria was noted by retrieving RlpA sequences, aligning them using
ClustalW2 (Larkin et al., 2007), and identifying what residue aligned with residue D168
of RlpA from P. aeruginosa. Multiple RlpA sequences were obtained from the various
classes of proteobacteria. Deltaproteobacteria: Desulfarculus baarsii DSM 2075,
Desulfovibrio vulgaris Hildenborough, Pelobacter carbinolicus DSM 2380, and
Syntrophobacter fumaroxidans MPOB. Epsilonproteobacteria: Arcobacter butzleri ED-
41
1, Campylobacter jejuni jejuni NCTC 11168, Helicobacter pylori Cuz20, and Nautilia
profundicola. Alphaproteobacteria: Acetobacter pasteurianus pasteurianus IFO 3283-01,
Caulobacter crescentus CB15, Parvularcula bermudensis, and Phenylobacterium
zucineum HLK1. Betaproteobacteria: Bordetella pertussis CS, Gallionella
capsiferriformans, Neisseria gonorrhoeae FA, and Thiobacillus denitrificans ATCC.
Gammaproteobacteria: Enterobacter cloacae cloacae ATCC, Escherichia coli K-12
substr. MG1655, Klebsiella pneumoniae subsp. rhinoscleromatis, Pseudomonas
aeruginosa UCBPP-PA14, Salmonella enterica, Serratia marcescens FG194, Vibrio
parahaemolyticus RIMD, and Yersinia pestis A1122. Note that all RlpA sequences
contain a DPBB and SPOR domain as predicted by the Pfam database (Finn et al., 2014).
42
Table 2.1. Strains used in this study
Strain
Relevant features
Source or reference
BL21
dcm ompT hsdS(rB- mB-) gal [malB+]K-12(λS)
Lab collection
EC251
K-12 wild type MG1655
Lab collection
EC2219
BL21/pDSW1132
This work
EC2292
BL21(λDE3)/pET28a-AmiD
Tom Bernhardt
EC3087
BL21/pDSW1557
E. coli
EC3129
This work
q
MC4100 ∆(λattL-lom)::kan lacI P207::gfp-
This work
ftsI/pDSW930
EC3183
∆rlpA
This work
EC3204
BL21/pDSW1600
This work
EC3220
BL21/pDSW1601
This work
EC3223
BL21/pDSW1604
This work
EC3225
BL21/pDSW1606
This work
EC3433
∆amiA
This work
EC3437
∆amiC
This work
EC3439
∆rlpA ∆amiA
This work
EC3443
∆rlpA ∆amiC
This work
EC3486
∆amiA ∆amiC
This work
EC3492
∆rlpA ∆amiA ∆amiC
This work
EC3552
BL21/pDSW1674
This work
EC3702
∆mltA ∆mltD ∆mltE ∆slt ∆mltC::kan
This work
EC3704
∆rlpA ∆mltA ∆mltD ∆mltE ∆slt ∆mltC::kan
This work
EC3745
∆mltA ∆mltD ∆mltC::kan
This work
EC3747
∆rlpA ∆mltA ∆mltD ∆mltC::kan
This work
SM10
thi thr leu tonA lacY supE recA::RP4-2-Tc::Mu (Simon, 1983)
KanR
P. aeruginosa
MJ1
UCBPP-PA14 pathogenic isolate wild type
Lab collection
43
Table 2.1. continued
Strain
Relevant features
Source or reference
MJ7
PA14 rlpA::MAR2xT7
(Liberati et al., 2006)
MJ18
PA14 dacC::MAR2xT7
(Liberati et al., 2006)
MJ24
MJ1 ∆rlpA
This work
MJ26
MJ1 ∆rlpA/pJN105
This work
MJ27
MJ1 ΔrlpA/pDSW1398
This work
MJ34
MJ1 Δsltb1
This work
MJ36
MJ1 rlpA-mCherry
This work
MJ40
MJ1 ∆rlpA/pDSW1518
This work
MJ42
MJ1 rlpA(Δ269-341)-mCherry
This work
MJ47
MJ1 Δmltb1
This work
MJ49
MJ1 Δsltb1 Δmltb1
This work
MJ71
MJ1 ∆rlpA/pDSW1545
This work
MJ73
MJ1 ∆rlpA/pDSW1554
This work
MJ81
MJ1 rlpA(E120A)-mCherry
This work
MJ83
MJ1 rlpA(D123A)-mCherry
This work
MJ85
MJ1 rlpA(H131A)-mCherry
This work
MJ89
MJ1 rlpA(D157N)-mCherry
This work
MJ117
MJ1 ∆rlpA/pDSW1635
This work
MJ119
MJ1/pDSW1635
This work
MJ131
MJ1 ∆rlpA/pDSW1676
This work
MJ133
MJ1 ∆rlpA/pDSW1694
This work
MJ137
MJ1 ∆rlpA/pMRP9-1
This work
MJ138
MJ1 ∆rlpA/pDSW1695
This work
44
Table 2.2. Plasmids used in this study
Plasmid
Relevant features
Source or reference
pDSW912
P204 rfp fusion vector; AmpR lacIq pBR ori
Kyle Williams
pDSW913
P206 rfp fusion vector; AmpR lacIq pBR ori
(Arends et al., 2010)
pDSW930
pDSW912::rlpAEc-mCherry
David Weiss
pDSW1132
pQE-80L::rlpAEc(25-362)a
This work
pDSW1385
pEXG2::‘sltb1 ΔrlpA dacC’
This work
pDSW1398
pJN105::rlpAPa
This work
pDSW1399
pDSW913::rlpAPa-mCherry
This work
pDSW1489
pEXG2::‘rlpA-mCherry dacC’
This work
pDSW1490
pEXG2::‘rodA Δsltb1 rlpA’
This work
pDSW1497
pDSW913::rlpAPa(Δ269-341)-mCherry
This work
pDSW1504
pEXG2::‘rlpA(Δ269-341)-mCherry dacC’
This work
pDSW1516
pEXG2::‘PA14_57740 Δmltb1 cysD’
This work
pDSW1518
pJN105::rlpAPa-mCherry
This work
pDSW1519
pJN105::rlpAPa(E120A)-mCherry
This work
pDSW1520
pJN105::rlpAPa(D123A)-mCherry
This work
pDSW1537
pJN105::rlpAPa(H131A)-mCherry
This work
pDSW1545
pJN105::rlpAPa(D157N)-mCherry
This work
pDSW1554
pJN105::rlpAEc-mCherry
This work
pDSW1557
pQE-80L::rlpAPa(28-341)
This work
pDSW1600
pQE-80L::rlpAPa(D157N) (28-341)b
This work
pDSW1601
pQE-80L::rlpAPa(E120A) (28-341)b
This work
pDSW1604
pQE-80L::rlpAPa(D123A) (28-341)b
This work
pDSW1606
b
pQE-80L::rlpAPa(H131A) (28-341)
This work
pDSW1614
pEXG2::‘sltb1 rlpA-mCherry dacC’
This work
pDSW1615
pEXG2::‘sltb1 rlpA(E120A)-mCherry dacC’
This work
pDSW1616
pEXG2::‘sltb1 rlpA(D123A)-mCherry dacC’
This work
pDSW1617
pEXG2::‘sltb1 rlpA(H131A)-mCherry dacC’
This work
45
Table 2.2. continued
Plasmid
Relevant features
Source or reference
pDSW1619
pEXG2::‘sltb1 rlpA(D157N)-mCherry
This work
dacC’
pDSW1635
pJN105::amiB-mCherry
This work
a
pDSW1674
pQE-80L::rlpAEc(S147D) (25-362)
This work
pDSW1676
pJN105::rlpAPa(D168N)-mCherry
This work
pDSW1694
pJN105::rlpAPa(D168S)-mCherry
This work
pDSW1695
pJN105::rlpAEc(S147D)-mCherry
This work
pEXG2
Suicide vector; ColEI ori mob sacB GentR
(Rietsch et al., 2005)
pET28a-AmiD
his6-amiD
(Uehara & Park, 2007)
pJN105
Arabinose regulation (PBAD); pBBR ori
(Newman & Fuqua, 1999)
GentR
pMRP9-1
Constitutive expression of gfp in P.
(Davies et al., 1998)
aeruginosa; CarbR
pQE-80L
PT5 containing lac operators; lacIq ColE1
Qiagen
ori AmpR
a
The numbers 25-362 refer to the residues of RlpAEc included in the construct; the
construct removes the signal sequence (residues 1-17) and an additional 7 residues of
RlpAEc.
b
The numbers 28-341 refer to the residues of RlpAPa included in the construct; the first 27
a.a. of RlpAPa encode the signal sequence and were omitted.
46
Table 2.3. Primers used in this study
Primer
Sequencea
P1128
GCCAAGCTTTACTGCGCGGTAGTAATAAAT
P1140
CAGGAATTCATGCGTAAGCAGTGGCTCGGGA
P1141
GTCTCTAGAGTTGTTGTTCTGCGCGGTAGTAATAAATGAC
P1375
CAAAGATCTAACAACAACCAACAGACGGTAAGTGTA
P1474
AAAATCTAGAGGAGGAGCGGACACGCTTGCTC
P1475
AAAATCTAGACCGACGCTGGTACGCCCCGACTG
P1507
AAAAAAGCTTCGGCCCAGGCGGGGGACTAC
P1508
AAAACAATTGCTTCCAGACCAGGCCCTTGG
P1599
GCCGAATTCAGCAAGCGTGTCCGCTCCTCC
P1600
CTGTCTAGAGTTGTTGTTGTCGGGGCGTACCAGCGTCGG
P1603
GCAGAATTCGACCAGAAGGTCACGGCGATG
P1604
CAATCTAGATCAGTCGGGGCGTACC
P1680
GCAAAGCTTAAGCGTGTCCGCTCCTCCCTG
P1681
GCCCAATTGTTACTTGTACAGCTCGTCCAT
P1682
GCACAATTGGCGCCTACTCACGCAGGGAAT
P1683
GGCGGTACCGTCATGGTCAGGTCTTCGGCG
P1702
CAGAAGCTTCATGCTGATGAAGCAGGCCAC
P1703
CTGCTCGAGCAGTACTTGCATTGCGTTCTT
P1704
CAGCTCGAGCGCGCGCGAGGTGCCCATTGA
P1705
CTGGAATTCTGCTGGTTGCGTACGACCGAG
P1708
CTGTCTAGAGTTGTTGTTGAGATACAGGCCATCGGCTGG
P1711
CTGAAGCTTCAGTCGGGGCGTACCAGCGTC
P1713
CAGAAGCTTGAAGGCAGCGTCGAAACCGTAC
P1714
CTGCTCGAGCAGGGCGAGGGCGGTACGGCG
P1715
CAGCTCGAGTCCGTCGTCAGGCAGGATTAG
P1716
CTGGGTACCCTGAGCACCCTGGTCGAAGAG
P1727
CTGGAGCTCTTACTTGTACAGCTCGTCCATG
P1754
TGGGACGTCGACGTGTCGCGGATC
47
Table 2.3. continued
Primer Sequencea
a
P1755
CATTCTAGAGTTGTTGTTGTCGGG
P1756
TAGAGGTCGTAGGTCGCGCCGTTGGCGGTGG
P1758
GTCATGCCGTAGAGGGCGTAGGTCTCGCCGT
P1760
AACGGCAGGGTCTTGGCCGCGGCGGTCATGCC
P1770
AGCGAATTCGACCAGAAGGTCACG
P1773
GCTGACCATCATCGCTTGTACAACTGCTCAACAGCACGGCCGC
P1774
CATTCTAGAGTTGTTGTTCTGC
P1781
ATAGAACGGGCCGCGGTTGTTGACGCGGACGATC
P1787
CATTGATCATCCAGCAAGGCGCCCCAGCAG
P1805
CAGGAATTCCCACCCTGACCATGGGAGCATG
P1806
CTGTCTAGAGTTGTTGTTCTGGGCCGCCAGGGCGGTGCT
P1821
CAGAAGCTTTACTGCGTACATGGGCGGCCAG
P1822
CGGGGATCCGCGACACGTCGACGTC
P1823
CGCGGATCCCCGATGCGGTGCCGA
P1824
ACGGCGGCCGCGTGCTGCGCCGGC
P1951
GTCAGCTGCCGCGCGAGAAAGGTCAATAACGCGGTCGTTGCCGTA
P1957
CTTCGCCGCGGCGAAGGACAGGTTGATGACCCGGTCGGAATAGAA
P1959
CTTCGCCGCGGCGAAGGACAGGGAGATGACCCGGTCGGAATAGAA
P1967
TAGCGAATTCGACCAGAAGGT
P1968
GACACTGAACTTGTTCCCGCG
All primer sequences are written 5’ to 3’. Restriction sites are underlined.
48
CHAPTER 3: THE BACTERIAL SEPTAL RING PROTEIN RLPA IS A
LYTIC TRANSGLYCOSYLASE THAT CONTRIBUTES TO ROD SHAPE
AND DAUGHTER CELL SEPARATION IN PSEUDOMONAS AERUGINOSA
Introduction
Most bacteria have a peptidoglycan (PG) cell wall that protects the organism from
lysis due to turgor pressure and confers on the cell its characteristic shape [reviewed in
(Vollmer et al., 2008a)]. The PG sacculus contains a carbohydrate backbone composed
of a repeating disaccharide of N-acetylglucosamine and N-acetylmuramic acid,
abbreviated here as NAG and NAM, respectively (Figure 3.1). These glycan strands are
cross-linked by oligopeptides attached to the NAM moieties. Because the sacculus is a
single, covalently-closed molecule that completely surrounds the cell, it must be
continually remodeled during growth and cell division. In particular, for rod-shaped
bacteria to elongate, PG in the lateral wall must be selectively hydrolyzed to make room
for insertion of new material, and selective hydrolysis of septal PG is required for
daughter cells to separate after cell division.
Bacteria typically produce multiple, seemingly redundant PG hydrolases that are
usually classified by the type of bond they cleave in PG (Figure 3.1A) (van Heijenoort,
2011, Vollmer et al., 2008b). These enzymes include amidases that liberate the stem
peptides from the glycan backbone, lytic transglycosylases that degrade the glycan
backbone, endopeptidases that cleave cross-links between adjacent stem peptides, and
carboxypeptidases that trim the ends of stem peptides (Figure 3.1A). Whereas the
enzymatic activity of the various PG hydrolases is usually clear-cut, their precise
physiological roles are often difficult to establish because mutants lacking one or two of
these enzymes frequently grow and divide normally. But at least in E. coli, mutants
49
lacking larger numbers of PG hydrolases exhibit complex morphological abnormalities
(Heidrich et al., 2001, Heidrich et al., 2002, Potluri et al., 2012, Priyadarshini et al.,
2007). These observations point to extensive functional redundancy and suggest some
hydrolases contribute to both elongation and daughter cell separation. Nevertheless,
studies in E. coli have highlighted the importance of amidases for daughter cell
separation and endopeptidases for elongation (Heidrich et al., 2001, Priyadarshini et al.,
2007, Uehara et al., 2010, Singh et al., 2012). Endopeptidases also play a critical role in
elongation in Bacillus subtilis and Vibrio cholerae (Hashimoto et al., 2012, Dörr et al.,
2013).
The focus of this manuscript is an outer membrane lipoprotein of previously
unknown function named RlpA (rare lipoprotein A), which we show below is an unusual
lytic transglycosylase—it preferentially digests “naked” glycan strands that lack stem
peptides. Prior to this report, RlpA had only been studied in E. coli, where several
observations linked the protein to morphogenesis and peptidoglycan metabolism, albeit
only in indirect ways. Fusions of mCherry to RlpA revealed localization to scattered foci
along the lateral wall (Gerding et al., 2009) and, even more prominently, to the septal
ring that mediates cell division (Gerding et al., 2009, Arends et al., 2010). In E. coli,
rlpA is in an operon with pbpA and rodA, which encode proteins needed for
peptidoglycan synthesis during elongation (Figure 3.2A) (Matsuzawa et al., 1989,
Mohammadi et al., 2011, Banzhaf et al., 2012). Immediately downstream but
transcribed separately is dacA, which codes for a peptidoglycan hydrolase implicated in
spatial control of cell division (Figure 3.2A) (Potluri et al., 2012). The sequence of RlpA
contains two domains, a C-terminal “SPOR domain” (Pfam 05036) and a central “RlpA-
50
like double-psi beta-barrel domain” (DPBB; Pfam 03330) (Figure 3.2B) (Punta et al.,
2012). SPOR domains are about 75 amino acids long, bind peptidoglycan, and localize
to the septal ring (Ursinus et al., 2004, Möll & Thanbichler, 2009, Gerding et al., 2009,
Arends et al., 2010). Most characterized SPOR domain proteins are involved in cell
division, although at least two are involved in other aspects of morphogenesis (Mishima
et al., 2005, Gode-Potratz et al., 2011). DPBB folds are found in many proteins and are
often enzymatic, but the activity is different in different proteins [reviewed in (Castillo et
al., 1999)]. Threading the DPBB domain from E. coli RlpA by the Protein
Homology/analogY Recognition Engine (PHYRE) (Kelley & Sternberg, 2009) revealed
distant similarity to expansin-like cellulose binding domains, which bind carbohydrates
but are not enzymatic (Sampedro & Cosgrove, 2005), and to the MltA lytic
transglycosylase of E. coli (van Straaten et al., 2005). Neither of these similarities is
strong enough to be detected in a BLAST search.
Collectively, these observations suggest RlpA might be an enzyme involved in
synthesis or degradation of PG during division and/or elongation, but E. coli null mutants
of rlpA do not have any obvious morphological defects (Gerding et al., 2009, Arends et
al., 2010). Moreover, in our hands purified RlpA from E. coli does not digest PG sacculi
isolated from wild-type cells. What broke this impasse was the fortuitous observation
that in P. aeruginosa an rlpA null mutant has striking morphological defects that link the
protein to division and rod shape. Follow-up studies revealed P. aeruginosa RlpA is a
lytic transglycosylase whose activity appears to be restricted to “naked” glycan strands
that lack stem peptides.
51
Results
An rlpA mutant has a chaining phenotype in P. aeruginosa. RlpA appears to
be the most highly conserved of all the SPOR domain proteins, with over 5000 examples
from more than 2,500 species listed in the Pfam database (version 27.0) (Punta et al.,
2012). Conservation is usually a hallmark of importance, yet of the four SPOR domain
proteins in E. coli, RlpA is the only one that appears to be completely dispensable
(Gerding et al., 2009, Arends et al., 2010). We therefore decided to analyze RlpA in
other bacterial species in hopes of finding a useful phenotype. Utilizing the BLAST
function on the Pseudomonas Genome Database website (Winsor et al., 2011), we
identified rlpA in strain PA14 as PA14_12090. The E-value for comparison of the E. coli
and P. aeruginosa RlpA proteins is 10-24. The two proteins are very similar in overall
size and domain structure (Figure 3.2B). In both organisms rlpA appears to be
cotranscribed with genes involved in biogenesis of the PG sacculus (Figure 3.2A), but
there is one striking difference— the gene immediately upstream of rlpA in P. aeruginosa
encodes a soluble lytic transglycosylase designated sltb1that is not found in the E. coli
operon (Blackburn & Clarke, 2002, Nikolaidis et al., 2012).
The PA14 transposon insertion library contains a single insertion mutation of rlpA
(rlpA::Tn); the insertion site is 138 base pairs downstream of the first T in the TTG start
codon (http://ausubellab.mgh.harvard.edu/cgi-bin/pa14/home.cgi) [11, December 2013]
(Liberati et al., 2006). We obtained the rlpA::Tn mutant, confirmed the insertion site by
PCR, and tested its phenotypes under various growth conditions. The mutant grew
normally on LB plates containing 10 g/L NaCl, but, to our surprise, was not viable when
plated on LB lacking NaCl (hereafter designated LB0N) over a range of temperatures
52
(shown for 37°C in Figure 3.2C). The rlpA::Tn mutant appeared normal in LB broth, but
in LB0N it grew slowly and formed chains of unseparated cells (Figure 3.2D). These
phenotypes did not result from polarity onto dacC because a dacC::Tn mutant (from the
same mariner insertion library) had a similar plating efficiency to wild type on LB0N
plates and the cells looked normal when grown in LB0N broth (Figure 3.2C and 3.2D).
We then constructed an in-frame deletion of rlpA, which phenocopied the
rlpA::Tn mutant. Specifically, the ΔrlpA mutant was indistinguishable from wild type
when grown in LB broth (Figure 3.2E), but growth arrested about 2.5 hours after shift to
LB0N broth (Figure 3.2F) and the mutant failed to form colonies when plated on LB0N
(Figure 3.2C). Microscopy of cells grown in LB0N broth confirmed a chaining defect,
which became more pronounced the longer the cultures were allowed to grow (Figure
3.2D and 3.3B). Close inspection of cells in the chains revealed they were ~50% shorter
and 20% wider than wild type (Table 3.1). Analysis of cells in the chains by fluorescence
loss in photobleaching (FLIP) revealed 84% of the septa were closed, indicating that
membrane constriction had gone to completion (Figure 3.4A). The morphological and
viability defects could be rescued by expressing rlpA from a plasmid (Figure 3.2C and
3.2D). The mutant could also be rescued by replacing NaCl in the growth medium with
proline or sucrose (Figure 3.4B), indicating the phenotypic changes are due to a general
osmotic stress rather than specifically related to NaCl. Time-lapse microscopy of live
cells in LB0N spotted on an agarose pad revealed about half of the cells lysed, while the
other half stopped growing but remained phase dark (Figure 3.4C). Collectively, our
findings demonstrate that RlpA is important for daughter cell separation and rod shape
when P. aeruginosa is grown in medium of low osmotic strength.
53
Septal localization of P. aeruginosa RlpA. To explore localization of the P.
aeruginosa protein, we replaced the chromosomal rlpA allele with an rlpA-mCherry gene
fusion. Western blotting with anti-mCherry sera indicated the fusion protein was stable
(Figure 3.5A). The RlpA-mCherry fusion protein was functional as evidenced by
viability on LB0N plates (Figure 3.5B) and absence of chaining in LB0N broth (Figure
3.6A-C; Table 3.2). Fluorescence microscopy of live cells grown to midlog phase in LB
revealed septal localization in ~42% of the cells in the population (n > 500 cells; Figure
3.4D). Most of these cells had obvious constrictions, suggesting RlpA is a late recruit to
the septal ring. Polar localization was observed in ~15% of the cells (Figure 3.4D).
Because most of these cells were short, we suspect this reflects persistence of RlpAmCherry after division is complete. Finally, we observed weak foci along the lateral wall
in ~5% of the cells, which might reflect a role for RlpA in elongation, peptidoglycan
recycling or tailoring of the lateral wall. In total, the localization patterns seen in P.
aeruginosa are similar to what has been reported in E. coli (Gerding et al., 2009, Arends
et al., 2010).
To determine if the SPOR domain is needed to target RlpA to the midcell, we
replaced the chromosomal allele of rlpA with an rlpA(ΔSPOR)-mCherry construct. The
fusion protein was stably produced (Figure 3.5A) and functional (Figure 3.5B; Table 3.2),
but septal localization was barely detectable and foci were no longer observed along the
lateral wall (Figure 3.6D). Thus, the SPOR domain of RlpA is very important for normal
localization, but not for cell division or rod shape. This paradoxical situation has been
reported previously for FtsN, which is targeted to the division site by its SPOR domain
but nevertheless supports cell division even after the SPOR domain has been deleted
54
(Ursinus et al., 2004, Möll & Thanbichler, 2009, Gerding et al., 2009). The most
plausible interpretation is that small amounts of properly localized RlpA are sufficient for
biological function.
Low osmolarity does not induce rlpA. Because the rlpA-mCherry fusion was
integrated into the native locus, we used it to ask whether rlpA expression is osmoregulated. Essentially identical steady-state levels of RlpA-mCherry protein were
detected by Western blotting with anti-mCherry when cells were grown in LB or LB0N
(Figure 3.7). Thus, although osmotic stress is needed to uncover the phenotypes
associated with loss of rlpA, the gene does not appear to be part of an osmotic stress
response regulon.
PG from the ΔrlpA mutant is enriched in naked glycans. The chaining
phenotype suggested RlpA is a PG hydrolase, but we were unable to detect hydrolase
activity when purified RlpA was incubated with sacculi isolated from wild-type cells,
despite exploring a number of assay formats (documented below). We therefore turned
to analysis of the PG in hopes of identifying structural abnormalities that would provide
some insight into what RlpA does. For these experiments wild type and the ΔrlpA
mutant were grown for several generations in LB0N until the mutant had formed chains
4-8 cells in length. Comparison of the HPLC elution profiles of muropeptides obtained
after muramidase digestion of sacculi revealed several differences, the most striking of
which were that the mutant was enriched in three muropeptides that eluted from the
HPLC column with retention times of 15 min (P5), 23 min (P9) and 29 min (P13) (Figure
3.8, Table 3.3). The peaks were identified using a combination of amino sugar analysis,
amino acid analysis and mass spectrometry methods (Table 3.4 and 3.5).
55
The species eluting as P5 contained abundant NAG and NAM but little in the way
of amino acids, suggesting it is a fragment of the glycan backbone that lacks peptide side
chains. The P5 product had a mass-charge ratio (m/z) of 999.4 Da, consistent with it
being the tetrasaccharide NAG-NAM-NAG-NAMol (abbreviated TS, predicted m/z =
999.5 Da; the terminal NAM is in the alcohol form due to borohydride reduction). This
was confirmed by fragmentation analysis with tandem mass spectrometry (Table 3.5).
The P9 product was also highly enriched in amino sugars as compared to amino acids,
and had a mass-charge ratio of 1477.6 Da, consistent with the hexasaccharide NAGNAM-NAG-NAM-NAG-NAMol (abbreviated HS, predicted m/z = 1477.6). Finally, the
P13 product had excess amino sugars as compared to amino acids, but the ratio was not
as skewed as for the other two products. The mass-charge ratio was 1442.9, consistent
with a tetrasaccharide that contains one tetrapeptide side-chain: NAG-NAM-NAGNAMol-Ala-Glu-Dpm-Ala (abbreviated TS-Tetra, predicted m/z = 1442.6 Da). This
structure for the P13 product was consistent with at least 9 fragments in tandem mass
spectrometry (Table 3.5). The fact that we recovered tetra- and hexasaccharide fragments
indicates mutanolysin does not cleave very efficiently at NAM residues that lack stem
peptides. A lack of mutanolysin digestion adjacent to a peptide-free NAM within a
tetrasaccharide was previously observed (Gilmore et al., 2004). Mutanolysin also fails to
cut at muramic δ–lactam (Popham et al., 1996a), a modified form of NAM that also lacks
a stem peptide and is abundant in bacterial spore PG (Warth & Strominger, 1969). Thus,
when doing muropeptide analysis of PG it is not safe to assume that all fragments have
been reduced to disaccharides.
56
Besides the above-mentioned muropeptides that were more abundant in the
mutant, two muropeptides were less abundant (Figure 3.8, Table 3.3). These eluted at 14
min (P4) and 22 min (P8), and proved to be disaccharides with a tripeptide or
pentapeptide sidechains, respectively. We do not know the significance of these changes,
which in any event were small compared to the increases in the P5, P9 and P13
muropeptides.
In summary, PG from the ΔrlpA mutant accumulated regions of glycan strand that
lack stem peptides. This finding suggested RlpA degrades naked glycans and explained
why RlpA failed to exhibit hydrolase activity in our in vitro assays, because sacculi from
wild-type cells are essentially devoid of naked glycans (Figure 3.8, Table 3.3).
RlpA is an unusual lytic endo-transglycosylase with specificity for glycan
strands that lack stem peptides. For enzymological assays we purified a soluble
derivative of RlpA that carried a hexahistidine tag in place of the N-terminal type II
signal sequence (His6-RlpA; Figure 3.9A). As substrate we prepared PG sacculi that had
been labeled with the dye Remazol Brilliant Blue R (RBB-PG) (Zhou et al., 1988).
Although His6-RlpA did not have convincing hydrolase activity when incubated with
RBB-PG sacculi from wild-type cells (as expected), it readily hydrolyzed sacculi from
the ΔrlpA mutant (Figure 3.9B and C). This was confirmed using muropeptide analysis
to examine the effect of incubating His6-RlpA with unlabeled sacculi. In the case of
sacculi from wild-type cells, no PG fragments were released into the soluble fraction, and
analysis of the residual insoluble pellet indicated it was not changed relative to the
starting material (Figure 3.10). But when His6-RlpA was incubated with sacculi from the
rlpA mutant, P5 (tetrasaccharide) and to a lesser extent P9 (hexasaccharide) disappeared
57
from the insoluble fraction and two products appeared in the supernatant (Figure 3.9D
and 3.10). These products eluted from the HPLC column at 33 min (Peak a) and 43 min
(Peak b). The first product was identified by mass spectrometry as a tetrasaccharide with
a 1,6-anhydro end: NAG-NAM-NAG-1,6-anhydroNAM (observed m/z = 979.3 Da,
predicted = 979.4 Da). Similarly, the second product was identified as a hexasaccharide
with a 1,6-anhydro end: NAG-NAM-NAG-NAM-NAG-1,6-anhydroNAM (observed m/z
= 1457.8 Da, predicted = 1457.5 Da). These identifications were confirmed by the
observation of appropriate fragmentation products during tandem mass spectrometry
(Table 3.6). Moreover, the HPLC retention time was not affected by treatment with the
reducing agent NaBH4 (Figure 3.9D), consistent with the presence of a non-reducible 1,6anhydroNAM end. The presence of 1,6-anhydroNAM ends in the cleavage products
indicates RlpA is a lytic transglycosylase rather than a muramidase or glucosamidase
(Holtje et al., 1975, Vollmer et al., 2008b). The relatively large size of the released
products (tetra- and hexasaccharides) indicates RlpA is an “endo” enzyme that cuts in the
middle of glycan chains rather than an “exo” enzyme that cuts near the end of glycan
chains to release disaccharides. We purified sufficient P5 (tetrasaccharide) to use as a
substrate for His6-RlpA, but no hydrolysis was detected, suggesting RlpA requires greater
context before cleaving (Figure 3.11).
Several aspects of the assays shown in Figure 3.9 need to be clarified to prevent
potential points of confusion. The reason His6-RlpA can almost completely solubilize
dye-labeled sacculi in Figure 3.9C despite having very limited substrate specificity is that
much of the “soluble” PG is in the form of very large fragments that remain in the
supernatant when samples are centrifuged for 10 min to pellet residual insoluble sacculi.
58
This also accounts the apparent discrepancy between extensive solubilization on the one
hand and the recovery of only a small amount of soluble products after HPLC on the
other (Figure 3.9C vs. 3.9D). Most of the “soluble” PG fragments released during RlpA
treatment are too large for HPLC analysis, which in Figure 3.9D did not include a
muramidase digestion step. Consistent with this, if the soluble fraction is treated with
muramidase prior to loading onto the HPLC column, we observe more total material and
a variety of muropeptide structures (data not shown). Finally, the inference that RlpA is
lytic transglycosylase is only valid if the 1,6-anhydroNAM ends were introduced by
RlpA cleavage. Note that RlpA digestion of sacculi from the ΔrlpA mutant greatly
diminished the abundance of tetra- and hexasaccharides that lack 1,6-anhydroNAM ends
(Figure 3.9D, peaks 5 and 9) while at the same time releasing tetra- and hexasaccharides
that contain 1,6-anhdyroNAM ends (Figure 3.9D, peaks a and b). We take these findings
to mean ΔrlpA sacculi contained longer naked glycans that were cleaved by muramidase
to produce oligosaccharides with NAM ends or by RlpA to produce oligosaccharides
with 1,6-anhydroNAM ends.
RlpA degrades the product of amidase digestion. These results suggest RlpA
degrades the carbohydrate backbone of PG only after amidases have removed stem
peptides. This is unusual, but has been reported previously for SpoIID of Bacillus
subtilis (Morlot et al., 2010) and MltE of E. coli (Kraft et al., 1998), although some
reports indicate these enzymes also cleave glycans that have stem peptides under some
circumstances (Kraft et al., 1998, Gutierrez et al., 2010, Lee et al., 2013). In a recent
study of SpoIID (Morlot et al., 2010), the purified protein only degraded the glycan
backbone if the reaction mixtures included an amidase to remove stem peptides. We
59
used a modified version of that assay to show the same is true of RlpA. RBB-labeled
sacculi from wild type cells were subjected to a limited digestion with His6-AmiD from
E. coli, then the enzyme was inactivated by heating to 95°C. Upon addition of His6RlpA, robust dye release was observed; in four independent replicates AmiD treatment
stimulated RlpA activity by 5.7 ± 2.1 fold (mean ± standard deviation) (Figure 3.12A and
C, Figure 3.13). Conversely, pre-treatment of wild-type sacculi with His6-RlpA did not
stimulate subsequent hydrolysis by His6-AmiD (Figure 3.12B and C, Figure 3.13). The
fact that stimulation depended on the order of addition indicates dye release is not simply
a consequence of the cumulative activity of the two enzymes but instead means that
AmiD creates the substrate cleaved by RlpA.
Residue D157 in the DPBB is critical for lytic transglycosylase activity. We
used Clustal Omega (Sievers et al., 2011) to create a multiple sequence alignment of
DPBB domains from various RlpAs and identified highly conserved amino acids (Figure
3.14A). Based on this alignment, we constructed the following four mutant variants:
E120A, D123A, H131A and D157N. The mutant derivatives were purified and tested for
lytic transglycosylase activity in the dye release assay with ΔrlpA sacculi as substrate
(Figure 3.9A and C). Dye release activity was greatly reduced in every case, indicating
the targeted residues are important for lytic transglycosylase activity. In particular, the
D157N protein had no detectable enzymatic activity.
To assess the importance of these residues for RlpA function in vivo, the mutant
derivatives were fused to mCherry and recombined into the native rlpA locus. All of the
mutant proteins were produced in normal amounts and localized to the septal ring (Figure
3.5A, Figure 3.6E-H), but only the D157N lesion phenocopied the ΔrlpA mutant—it
60
failed to support growth on LB0N plates (Figure 3.5B) and microscopy revealed chains
of short, fat cells when grown in LB0N broth (Figure 3.6H and Table 3.2). The second
most defective protein in the dye release assay, the E120A mutant, resulted in slightly
increased chaining as compared to wild type (Table 3.2). Collectively, these findings
imply the enzymatic activity of RlpA is important for proper growth and division but low
levels of activity are sufficient. If the results from the dye-release assay can be taken at
face value, it appears that the break-point is around 6% of wild-type activity (i.e., the
activity of the E120A protein). This is consistent with the observation that the ΔSPOR
domain variant of RlpA supports proper growth and division even though it does not
localize very efficiently to the midcell (Figure 3.6D). It should be noted that P.
aeruginosa has 9 additional lytic transglycosylases, some of which might compensate for
lack of RlpA (Legaree & Clarke, 2008).
Evidence that RlpA is not needed for proper function of SltB1, MltB1 or
AmiB. Early on during this investigation we had observed that RlpA is needed for
efficient daughter cell separation, but purified RlpA did not appear to be a PG hydrolase,
so we invested some effort in in exploring whether RlpA activates or recruits a PG
hydrolase. Although this line of investigation was abandoned once we discovered that
RlpA had a lytic transglycosylase activity, we had by that time learned some new things
about three P. aeruginosa PG hydrolases: SltB1, MltB1 and AmiB.
We first investigated the lytic transglycosylases SltB1 and MltB1 (Blackburn &
Clarke, 2002, Scheurwater et al., 2007, Cavallari et al., 2013). These enzymes were
chosen because sltB1 is adjacent to rlpA (Figure 3.2A) and MltB1 is ~40% identical to
SltB1. We constructed in-frame deletions of sltB1 and mltB1, and observed that both
61
mutants exhibited normal morphology when grown in LB, as previously reported
(Blackburn & Clarke, 2002). More importantly for our purposes, morphology was also
normal in LB0N (Figure 3.15A). Even a ∆sltb1Δmltb1 double mutant appeared normal
in both media (Figure 3.15A). We conclude that the phenotypic changes cause by loss of
RlpA cannot be explained by failure to activate SltB1 and MtlB1.
We then turned our attention to amidases, because amidase mutants of E. coli
have chaining and shape defects similar to rlpA mutants of P. aeruginosa (Heidrich et al.,
2001, Priyadarshini et al., 2007). Moreover, AmiB and AmiC localize to the septal ring
in E. coli (Bernhardt & de Boer, 2003, Peters et al., 2011). Consistent with this, a P.
aeruginosa AmiB-mCherry fusion protein localized sharply to the septal ring, but this did
not require RlpA (Figure 3.15B). Therefore, the morphological defects observed in our
ΔrlpA mutant are not due to failure to recruit AmiB to the septal ring. Nevertheless,
AmiB is probably involved in daughter cell separation in P. aeruginosa and might work
together with RlpA. We also attempted to localize AmiA of P. aeruginosa but our
fusions to GFP and mCherry were retained in the cytoplasm (data not shown). It is likely
that AmiA localizes to septal regions because, in spite of its name, AmiA of P.
aeruginosa corresponds to the septal ring amidase AmiC of E. coli.
Discussion
Having failed to find a phenotype for a ΔrlpA mutation in E. coli, we turned to P.
aeruginosa. We chose this organism not because we had any reason to expect a different
outcome, but because an rlpA::Tn mutant was readily available (Liberati et al., 2006).
Interestingly, the rlpA::Tn mutant formed chains of short, fat cells when grown in LB
lacking NaCl. Follow-up studies revealed RlpA is a lytic transglycosylase and
62
contributes to both daughter cell separation and rod shape when P. aeruginosa is grown
in media of low osmotic strength. Further studies will be needed to determine why loss
of RlpA does not cause a similar phenotype in E. coli, but we anticipate RlpA will prove
to be important in many bacteria. In support of this, RlpA is well-conserved, with over
5,000 examples from more than 2,500 species in the Pfam database (Punta et al., 2012).
Moreover, Chaput et al. reported that a Tn insertion in mltD of Helicobacter pylori
caused a chaining phenotype, but an in-frame deletion of mltD did not (Chaput et al.,
2007). Intriguingly, the gene immediately downstream of mltD is rlpA.
RlpA is a new lytic transglycosylase with an unusual specificity for naked
glycans. Most characterized lytic transglycosylases solubilize whole PG sacculi because
they can efficiently cleave glycan strands containing stem peptides (Vollmer et al.,
2008b, Scheurwater et al., 2008, Lee et al., 2013). In contrast, in the case of RlpA we
only observe cleavage of glycan strands that lack peptide side-chains, although it should
be noted that a weak activity towards glycans that have stem peptides cannot be excluded
at this point because we do not know the detection limits of our assays. The substrate
preference of RlpA implies it digests glycan strands that have already been processed by
cell wall amidases, supporting the view that amidases are the pace-makers for cell
separation (Heidrich et al., 2001, Uehara et al., 2010, Uehara & Bernhardt, 2011). The
molecular basis for RlpA’s unusual specificity will require further investigation. It could
arise from the SPOR domain, which probably binds naked glycan strands (Ursinus et al.,
2004, Gerding et al., 2009). Alternatively, or in addition, the enzymatic domain may be
highly specific. This would account for the ability of RlpA to support normal growth and
63
division even after the SPOR domain is deleted (Figure 3.6D) and would also explain
why RlpA does not lyse cells when overproduced (data not shown).
The specificity of RlpA has implications for two long-standing questions about
PG hydrolases in general (Vollmer et al., 2008b, Uehara & Bernhardt, 2011). The first
question is, how do the various PG hydrolases work together to facilitate growth and
division? Our findings indicate RlpA and the amidases cleave PG in an ordered and
sequential fashion—first the amidases remove stem peptides, then RlpA degrades the
residual glycan backbone (Figure 3.1B). The second general question one must ask of all
PG hydrolases is what holds them in check so that they do not inadvertently cause lysis?
We think RlpA cannot lyse cells on account of its very limited substrate specificity.
Moreover, the activity of RlpA is regulated on at least two levels—by the SPOR domain,
which recruits the protein to the septal ring and by the amidases, whose activity is
regulated by a host of septal ring-associated proteins (Uehara et al., 2010, Yang et al.,
2011, Yang et al., 2012).
Models for how RlpA could facilitate daughter cell separation and
maintenance of rod shape. Cell wall amidases are widely considered the most
important enzymes for daughter cell separation, while the endopeptidases are generally
thought to be the key enzymes for elongation (Heidrich et al., 2001, Priyadarshini et al.,
2007, Hashimoto et al., 2012, Dörr et al., 2013, Singh et al., 2012). Nevertheless, our
study of RlpA is not the first to implicate lytic transglycosylases in these processes as
well. For example, an E. coli mutant lacking three lytic transglycosylases (ΔmltCDE) has
a mild chaining phenotype, while a mutant lacking six (ΔsltΔmltABCDE) has a stronger
chaining phenotype—and the cells become short and coccoid, indicating that lytic
64
transglycosylases are important not only for division but also for proper biogenesis of the
lateral wall (Heidrich et al., 2002). Chaining has also been reported for a ΔmltCΔmltE
double mutant in Salmonella enterica (Monteiro et al., 2011) and a single deletion of ltgC
(E. coli mltA homolog) in Neisseria gonorrhoeae (Cloud & Dillard, 2004). These
findings, together with the phenotypes reported here for ΔrlpA, suggest many lytic
transglycosylases play important roles in daughter cell separation and maintenance of cell
shape.
But why? Most current models of the PG sacculus indicate the glycan strands run
roughly perpendicular to the long axis of the cell (Vollmer & Höltje, 2004, Gan et al.,
2008), although a recent study indicates the glycan strands are not so highly organized
(Turner et al., 2013). In the standard model, it would seem amidase activity should be
sufficient for separation of daughter cells, so it is not obvious how a lytic
transglycosylase would help this process, all the more so in the case of RlpA, which
probably only digests PG that has already been cut by an amidase. We suggest that if the
division plane and glycan strands are not perfectly aligned, the glycan stands will cross
the division plane and be shared by daughter cells, leading to a situation in which both
amidases and lytic transglycosylases are needed for efficient daughter cell separation.
Likewise, irregularities in the organization of the PG might make efficient elongation
dependent upon removal of glycan strands rather than simply breaking crosslinks.
Alternatively, RlpA might affect cell separation and rod shape less directly by
contributing to PG recycling, tailoring the cell wall, or as part of a multiprotein complex
that does not function well when RlpA is missing or defective.
65
Comparison to MltE and SpoIID. Our findings bring to three the number of
unique lytic transglycosylases known to be exclusively or primarily active on glycan
strands that lack stem peptides: MltE of E. coli (Kraft et al., 1998), SpoIID of B. subtilis
(Morlot et al., 2010) and RlpA of P. aeruginosa. Curiously, these three proteins are not
homologous to one another and probably have completely different folds, yet they all
employ either a glutamate (MltE, SpoIID) or an aspartate (RlpA) as the general acid/base
during catalysis (Morlot et al., 2010, Artola-Recolons et al., 2011, Fibriansah et al.,
2012). Both MltE (Kraft et al., 1998) and RlpA (Figure 3.9D) are “endo” lytic
transglycosylases that cleave internal to glycan chains, whereas SpoIID is an “exo”
enzyme that releases disaccharides from the end of glycan (Morlot et al., 2010). SpoIID
is part of a protein complex required for the engulfment step of sporulation, and works
together with SpoIIP, a cell wall amidase that is part of the same complex (Abanes-De
Mello et al., 2002, Chastanet & Losick, 2007). Whether MltE in E. coli and RlpA in P.
aeruginosa have a dedicated amidase is not known, but AmiB is a likely partner for RlpA
in view of our finding that both proteins localize to the division site.
Potential new insights into MltA, a bacterial “expansin”, and a protein of
unknown function. Structural modeling using PHYRE (version 2.0) revealed the
catalytic DPBB domain of RlpA has intriguing similarity to several proteins in the
structure databases. One of these is the lytic transglycosylase MltA of E. coli, which has
been reported to be equally active on glycan strands with and without stem peptides
(Romeis et al., 1993, Ursinus & Höltje, 1994). The relatively high activity of MltA on
naked glycan strands is interesting in light of its distant structural relationship to RlpA,
which is specific for this substrate. The catalytic site of MltA contains two highly
66
conserved aspartates, D297 and D308, both of which are important for catalysis,
especially D308, which acts as the general acid/base (van Straaten et al., 2005, Powell et
al., 2006, van Straaten et al., 2007). These residues align with the conserved residues
D157 and D168 of P. aeruginosa RlpA, suggesting D168 is the catalytic acid/base
(Figure 3.14B and 3.16). But in E. coli RlpA this residue is a serine (Figure 3.14B), and
our data indicate D157 of the P. aeruginosa protein is important for lytic transglycosylase
activity (though we have yet to test D168). Further work will be needed to determine
what role the conserved aspartates play in RlpA and whether the E. coli protein has
enzyme activity. According to the PHYRE model of RlpA on MltA, none of the other
amino acids we targeted for mutagenesis in RlpA are in the catalytic pocket, although
some are close.
RlpA also showed similarity to YoaJ (EXLX1) from B. subtilis. YoaJ is reported
to be an expansin (Kerff et al., 2008), a class of proteins found mainly in plants.
Expansins are not catalytic but bind cellulose and loosen its structure (Sampedro &
Cosgrove, 2005). The report (Kerff et al., 2008) that concluded YoaJ is an expansin
considered the possibility that it is a PG hydrolase but ruled this out because (i) the
purified protein did not digest PG sacculi in vitro and (ii) a B. subtilis yoaJ null mutant
did not have an abnormal morphology. Interestingly, PHYRE models superimpose D71
of YoaJ with D157 of RlpA and D82 of YoaJ with D308 of MltA (Figure 3.14B). We
suggest YoaJ is a PG hydrolase, but like RlpA, it only digests naked glycan strands and is
only needed for proper morphology under a limited set of growth conditions.
The best match returned by PHYRE is to a protein of unknown function from P.
aeruginosa PAO1 designated PA4485 (Moynie et al., 2013). Similar to RlpA, PA4485 is
67
a predicted outer membrane lipoprotein that contains a DPBB fold, but unlike RlpA does
not contain a SPOR domain. Both catalytic aspartates (as inferred from RlpA and MltA)
are present in PA4485, suggesting it is a lytic transglycosylase. Neither the PAO1 nor
the PA14 transposon library (Jacobs et al., 2003, Liberati et al., 2006) contains
transposon insertions in PA4485, raising the possibility that PA4485 might be essential,
although this would be unprecedented for a lytic transglycosylase.
68
Table 3.1. Morphological parameters of a mutant lacking rlpA
Genotypea NaClb
Avg length
µm, (SD)
Avg unit
length µmc
Avg width
µm, (SD)
% Cells with indicated
no. of constrictions:
0
1
3
>3
+
2.9 (0.7)
2.3
0.9 (0.1)
74 26
0
0
3.1 (0.7)
2.3
0.9 (0.1)
73 27
0
0
+
3.1 (0.7)
2.4
0.9 (0.1)
75 25
0
0
ΔrlpA
5.9 (1.9)
1.1
1.1 (0.1)
0
5
66
29
a
Strains used were MJ1 (WT) and MJ24 (ΔrlpA). At least 300 cells were evaluated in
each case.
WT
b
c
Either 1% (+) or 0% (-) NaCl.
The distance between cell poles or constrictions in case of chains of cells.
69
Table 3.2. Functionality of various RlpA-mCherry fusion proteins
Genotypea
Avg length
µm, (SD)b
% Cells with indicated no. of
constrictions:
0
1
3
>3
rlpA::mCherry
3.1 (0.7)
74
26
0
0
ΔrlpA
5.8 (2.0)
0
10
60
30
rlpA(ΔSPOR)-mCherry
3.1 (0.7)
71
29
0
0
rlpA(E120A)-mCherry
3.5 (0.9)
55
42
3
0
rlpA(D123A)-mCherry
3.1 (0.7)
73
27
0
0
rlpA(H131A)-mCherry
3.1 (0.7)
71
29
0
0
rlpA(D157N)-mCherry
5.6 (1.9)
0
14
58
28
a
Strains shown are (in the order listed) MJ36, MJ24, MJ42, MJ81, MJ83, MJ85 and
MJ89. At least 300 cells were scored for each.
b
End-to-end length regardless of whether constrictions were observed.
70
Table 3.3. Muropeptide analysis of PG from
Pseudomonas aeruginosa PA14 WT and ΔrlpA grown in LB0N
Muropeptide Structureb
4
5
7
8
9
DS-Tri
TS
DS-Tetra
DS-Penta
HS
DS-TetraTetra
TS-Tetra
DS-TetraDS-Tetra
DS-TetraDS-Tetra
anhydro
11
13
17
20
% of all peaksc
Observed
m/z
[M+Na+]
893.5
999.5
964.5
1021.6
1477.7
Expected
m/z
[M+Na+]
893.4
999.3
964.4
1021.4
1478.4
1407.7
1407.6
1.9 (0.6) 1.7 (0.6)
1442.9
1443.4
1.0 (0.2) 1.9 (0.3)
1888.0
1887.8
27.1 (1.7) 25.1 (2.8)
1868.1
1867.8
4.9 (0.3) 6.4 (0.5)
WT
ΔrlpA
8.3 (0.2)
0.9 (0.6)
35.1 (1.2)
3.4 (0.3)
0.9 (0.6)
4.4 (1.8)
2.7 (0.7)
39.7 (2.4)
2.0 (0.3)
2.5 (1.2)
%
difference
-47.0
200.0
13.1
-41.2
177.8
-10.5
90.0
-7.4
30.6
a
Other
16.5
13.6
a
“Other” muropeptides includes the summed areas of multiple small peaks that did not
show significant variation between the two strains and that were not structurally
characterized.
b
Abbreviations: DS, disaccharide (NAG-NAMol); TS, tetrasaccharide (NAG-NAMNAG-NAMol); HS, hexasaccharide (NAG-NAM-NAG-NAM-NAG-NAMol). Tri,
tripeptide (L-Ala-D-iGlu-m-Dpm); Tetra, tetrapeptide (L-Ala-D-iGlu-m-Dpm-D-Ala);
Penta, pentapeptide (L-Ala-D-iGlu-m-Dpm-D-Ala-Gly). Anhydro, 1,6-anhydroNAM.
The terminal NAM is in the alcohol form due to borohydride reduction except in the case
of anhydro-NAM.
c
Percentages are the mean and standard error of three independent experiments.
71
Table 3.4. Amino acid and amino sugar analysis of muropeptides
Muropeptide
4
5
7
8
9
11
13
17
20
a
nmoles
NAG+NAMolb
4.3
2.8
14.6
1.3
1.2
0.6
0.6
12.1
1.7
Structure
Glu NAM
Ala
Dpm
DS-Tri
2.4
0.0
2.5
1.8
TS
0.4
1.3
0.1
0.1
DS-Tetra
7.7
0.0
17.4 7.0
DS-Penta
0.6
0.0
0.5
0.3
HS
0.0
0.8
0.0
0.0
DS-Tetra-Tetra
0.4
0.0
0.6
0.1
TS-Tetra
0.2
0.2
0.4
0.1
DS-Tetra-DS-Tetra 8.2
0.0
19.7 7.9
DS-Tetra-DS-Tetra 1.3
0.5
2.9
1.0
anhydro
a
Abbreviations: DS, disaccharide (NAG-NAMol); TS, tetrasaccharide (NAG-NAMNAG-NAMol); HS, hexasaccharide (NAG-NAM-NAG-NAM-NAG-NAMol). Tri,
tripeptide (L-Ala-D-iGlu-m-Dpm); Tetra, tetrapeptide (L-Ala-D-iGlu-m-Dpm-D-Ala);
Penta, pentapeptide (L-Ala-D-iGlu-m-Dpm-D-Ala-Gly). Anhydro, 1,6-anhydroNAM.
The terminal NAM is in the alcohol form due to borohydride reduction except in the case
of anhydro-NAM.
b
During analysis, NAMol and NAG co-elute, and NAMol standards produce
approximately 3-fold more absorbance than NAG standards, precluding a precise
quantification of each. Nanomole values in this table are calculated assuming a one-toone ratio of NAG and NAMol, which is true for disaccharides but not true for larger
muropeptides.
72
Table 3.5. Tandem mass spectrometry analysis of muropeptides P5, P9 and P13
Expected Observed
m/z
m/z
Formula
+
+
[M+Na] [M+Na]
a
C38H64N4O25
NAG-NAM-NAG-NAMol
999.4
999.5
b
C35H60N4O23
NAG-NAM-NAG-NAMol*
927.4
928.0
C30H51N3O20
NAM-NAG-NAMol
796.3
796.9
C27H44N3O17
NAG-NAM-NAG+c
704.3
704.8
C19H34N2O13
NAG-NAMol
521.2
521.6
+
C19H31N2O12
NAG-NAM
501.2
501.6
NAG-NAM-NAG-NAM-NAG-NAMol 1477.6
1477.6 C57H94N6O37
NAG-NAM-NAG-NAM-NAG1405.5
1406.0 C54H90N6O35
NAMol*
NAM-NAG-NAM-NAG-NAMol
1274.5
1275.0 C49H81N5O32
+
NAG-NAM-NAG-NAM-NAG
1183.4
1182.9 C46H74N5O29
C38H64N4O25
NAG-NAM-NAG-NAMol
999.4
999.7
NAG-NAM-NAG-NAM
980.4
979.7
C38H61N4O24
NAM-NAG-NAMol
796.3
796.6
C30H51N3O20
+
C27H44N3O17
NAG-NAM-NAG
704.3
704.5
C19H34N2O13
NAG-NAMol
521.2
521.4
C19H31N2O12
NAG-NAM+
501.2
501.4
NAG-NAM-NAG-NAMol-Ala-Glu1442.6
1442.9 C56H93N9O33
Dpm-Ala
NAG-NAM-NAG-NAMol-Ala-Glu1371.5
1371.1 C53H88N8O32
Dpm-Ala*
NAG-NAM-NAG-NAMol-Ala-Glu1354.5
1354.1 C53H87N8O31
Dpm
NAM-NAG-NAMol-Ala-Glu-Dpm-Ala 1239.5
1240.1 C48H80N8O28
NAG-NAM-NAG-NAMol-Ala-Glu
1182.5
1182.0 C46H76N6O28
NAG-NAM-NAG-NAMol-Ala
1070.4
1070.9 C41H69N5O26
NAG-NAMol-Ala-Glu-Dpm-Ala
964.4
964.8
C37H63N7O21
NAMol-Ala-Glu-Dpm-Ala
761.3
761.6
C29H50N6O16
+
C27H44N3O17
NAG-NAM-NAG
704.3
704.6
C19H31N2O12
NAG-NAM+
501.2
501.4
a
The terminal NAM is in the alcohol form due to borohydride reduction.
Structure
P13, Tetrasaccharide-tetrapeptide
P9, Hexasaccharide
P5,
Tetrasaccharide
Muropeptide
b
c
The asterisk indicates loss of a pyruval group from one of the NAM residues.
The + symbol (e.g., NAG-NAM+ or NAG-NAM-NAG+) indicates that a water molecule
has been lost during fragmentation.
73
Table 3.6. Tandem mass spectrometry analysis of muropeptides Pa and Pb
Pb, Hexasaccharide
anhydro
Pa, Tetrasaccharide
anhydro
Muropeptide
Expected
m/z
[M+Na]+
Observed
m/z
[M+Na]+
Formula
979.3
979.3
C38N4O24H60
907.3
907.2
C35N4O22H56
776.3
776.2
C30N3O19H47
NAG-NAM-NAG
704.3
704.2
C27N3O17H44
NAG-NAM anhydro
NAG-NAM-NAG-NAMNAG-NAM anhydro
NAG-NAM-NAG-NAMNAG-NAM anhydro*a
NAM-NAG-NAM-NAGNAM anhydro
NAG-NAM-NAG-NAMNAG+
NAG-NAM-NAG-NAM
anhydro
NAG-NAM-NAG-NAM
anhydro*a
NAM-NAG-NAM anhydro
501.2
501.1
C19N2O12H30
1457.5
1457.8
C57N6O36H90
1385.5
1385.2
C54N6O34H86
1254.5
1254.3
C49N5O31H77
1182.4
1182.3
C46N5O29H74
979.3
979.3
C38N4O24H60
907.3
907.3
C35N4O22H56
776.3
776.2
C30N3O19H47
704.3
704.2
C27N3O17H44
Structure
NAG-NAM-NAG-NAM
anhydro
NAG-NAM-NAG-NAM
anhydro*a
NAM-NAG-NAM anhydro
+b
+
NAG-NAM-NAG
a
NAG-NAM anhydro
501.2
501.1
C19N2O12H30
The asterisk indicates loss of a pyruval group from one of the NAM residues.
b
The + symbol (e.g., NAG-NAM+ or NAG-NAM-NAG+) indicates that a water molecule
has been lost during fragmentation. This explains why NAG-NAM+ and NAGNAManhydro have the same expected m/z ratio.
74
Figure 3.1. Model of PG and RlpA function. (A) Cartoon of the PG sacculus. Glycan
strands run roughly perpendicular to the long axis of the cell and are composed of a β1,4-linked NAG (G) and NAM (M). Short peptides (circles) are attached to the NAM
residues and cross-link the glycan strands. Lytic transglycosylases (LT) cleave the β-1,4
glucosidic bonds. Amidases (Ami) remove the stem peptides from the NAM residues.
Endopeptidases (EP) cleave the peptide cross-links. Carboxypeptidases (CP) remove
terminal amino acids from the stem peptides. (B) Model for sequential degradation of
PG by amidases and RlpA. First, amidases remove stem peptides from glycan strands.
Then RlpA cleaves the glycan strands, releasing mainly tetra- and hexasaccharides with a
1,6-anhydroNAM end (aM).
75
A
B
76
Figure 3.2. Growth and chaining of an rlpA mutant. (A) rlpA loci from E. coli K-12 and
P. aeruginosa UCBPP-PA14. Inverted triangles depict relative positions of the
MAR2xT7 transposon in rlpA and dacC. PBPA is a transpeptidase needed for crosslinking PG, especially during elongation. RodA is considered to be a flippase that
transports lipid-linked disaccharide-pentapeptide precursors to the periplasm for PG
synthesis during elongation. DacA is a PG carboxypeptidase. SltB1 is a soluble lytic
transglycosylase found in the periplasm. DacC is a PG carboxypeptidase more closely
related to DacC of E. coli than to DacA of E. coli. (B) Schematic diagram of the domain
architecture of RlpA. S, type II signal sequence. DPBB, RlpA-like double-psi betabarrel domain. SPOR, SPOR domain. DPBB domain residues targeted for mutagenesis
are shown below the P. aeruginosa protein. (C) Plating efficiency. Tenfold serial
dilutions of cells with the indicated genotypes were spotted onto LB (left) or LB0N
(right). Plates were photographed after incubation overnight at 37°C. P refers to the
empty vector (pJN105), while PrlpA refers to a derivative (pDSW1398) that carries rlpA.
(D) Division phenotypes. Cells grown at 37°C in LB or LB0N to an OD600 ~0.5 were
fixed, stained with the membrane dye FM4-64 and photographed under fluorescence.
The white bar represents 2 µm. (E, F) Growth curves for wild type and the ΔrlpA mutant
grown in LB or LB0N at 37°C. Strains shown are MJ1 (WT), MJ7 (rlpA::Tn), MJ18
(dacC::Tn), MJ27 (ΔrlpA/PrlpA), MJ26 (ΔrlpA/P) and MJ24 (ΔrlpA).
77
A
B
C
E
D
F
78
Figure 3.3. Scanning electron microscopy of a ΔrlpA mutant of P. aeruginosa. Wild
type strain MJ1(A) and ΔrlpA strain MJ24 (B) were grown at 37°C in LB0N from an
OD600 ~0.1 to an OD600 ~0.7, then fixed and prepared for SEM. The white bar represents
2 µm.
79
A
B
80
Figure 3.4. Phenotypes associated with rlpA. (A) The cytoplasm is compartmentalized
between cells in chains of a ΔrlpA mutant as demonstrated by fluorescence loss in
photobleaching (FLIP). The figure shows an overlay of DIC and fluorescence images of
strain MJ137, a ΔrlpA mutant harboring a plasmid that produces high levels of
cytoplasmic GFP. The cell to be bleached is indicated with an arrow. The cell was
bleached by iterative exposure to a beam of light from an argon laser. Cells were
photographed immediately before, immediately after, and 30 sec after bleaching. Note
that the neighboring cell did not lose fluorescence, indicating that septation had gone to
completion. A total of 21 cells from 16 different chains were analyzed by FLIP; those on
the end of a chain have only one septum but those internal to the chain have two septa, so
35 septa were tested in total. Of these, 30 were closed (86%) while 5 were open (14%).
(B) Rescue of ΔrlpA by osmolytes. Tenfold serial dilutions of WT and ΔrlpA cells were
spotted onto LB0N plates containing the indicated concentrations of NaCl, proline, or
sucrose. Plates were photographed after incubation overnight at 37°C. Strains shown are
MJ1 (WT) and MJ24 (ΔrlpA). (C) Two examples showing that the ΔrlpA mutant lyses
on LB0N. Numbers in the lower right refer to the time in minutes between images.
Strain MJ24 (ΔrlpA) in LB0N was spotted on an agarose pad and photographed under
phase contrast over a period of five hours. About 10% of the cells lysed during the
period of observation. In photographs taken the next morning, this had increased to 50%.
The remaining cells were phase-dark but did not grow after the first few hours. For the
cells that lysed, we observed a general disintegration of the wall and rounding-up, not
specific lysis at constrictions. Note that the cells shown here were maintained at room
temperature, whereas plates and growth curves shown elsewhere in this paper were
incubated at 37°C. Although the ΔrlpA mutant does not form colonies on LB0N plates at
room temperature, we do not know if the proportion of lysing cells is different at different
temperatures. (D) A field of cells showing representative results for localization of wildtype rlpA-mCherry. Filled arrows point to septal localization in cells at different stages
of the constriction process. Filled triangles point to examples of polar localization. Open
arrows point to foci along the lateral wall. The strain shown is MJ36 (rlpA-mCherry).
81
A
B
C
D
82
Figure 3.5. Phenotypes of rlpA mutants with a SPOR domain deletion or lesions in the
DPBB domain. (A) Western blot with anti-mCherry sera. Size markers are indicated to
the left of the blot. The predicted masses are 61 kDa for RlpA-mCherry and 53 kDa for
RlpA(ΔSPOR)-mCherry, assuming removal of the signal sequence. (B) Plating
efficiency, as in Figure 3.2C. The strains shown are MJ1 (WT), MJ36 (rlpA-mCherry),
MJ24 (ΔrlpA), MJ42 (ΔSPOR), MJ81 (E120A), MJ83 (D123A), MJ85 (H131A) and
MJ89 (D157N).
83
A
B
84
Figure 3.6. Function and localization of mutant derivatives of RlpA. Strains grown in
LB0N at 37°C to an OD600 ~0.5 were imaged by phase-contrast (left) and fluorescence
(right) microscopy. The fluorescence micrographs were inverted to better visualize
localization of mCherry fusion proteins. Arrows in (C) point to septal localization of
RlpA-mCherry. Chevrons in (D) point to sites with faint septal localization of
RlpA(ΔSPOR)-mCherry. The strains shown are listed in the legend to Figure 3.5.
85
86
Figure 3.7. RlpA is not upregulated by low osmolarity. Cells producing RlpA-mCherry
(strain MJ36) were grown to an OD600 ~0.5 in LB or LB0N before harvest. Whole-cell
extracts were diluted as indicated and subjected to Western blotting with anti-mCherry
sera. Molecular mass standards are shown at the left. The expected molecular mass of
RlpA-mCherry is 61 kDa after removal of the signal sequence.
87
88
Figure 3.8. ΔrlpA has PG alterations as compared to wild type. HPLC elution profiles of
muropeptides from WT (A) and ΔrlpA (B). PG sacculi were isolated from strains MJ1
(WT) and MJ24 (ΔrlpA) that had been grown to an OD600 ~0.5 in LB0N. Sacculi were
digested with mutanolysin and the resulting muropeptides were reduced with borohydride
prior to loading onto an RP-HPLC column. Muropeptide peaks are numbered and were
identified by amino acid and amino sugar analysis (Table 3.4) and tandem mass
spectrometry (Table 3.5). Arrows in (A) point to small peaks that are difficult to see.
89
A
B
90
Figure 3.9. RlpA is a lytic transglycosylase that cleaves naked glycan strands. (A)
Purified proteins (4 µg) were separated by SDS-PAGE (10% polyacrylamide) and stained
with Coomassie brilliant blue. WT = His6-RlpA. E120A, D123A, H131A, and D157N
are amino acid substitutions in RlpA. (B) Purified His6-RlpA does not solubilize dyelabeled PG sacculi from WT cells. Reaction mixtures contained 4 μM protein and were
incubated 18 hours at 37°C. Lysozyme (LZ) served as a positive control. Data shown
are mean and standard deviation of a representative experiment done in triplicate. All
experiments were done on at least 3 occasions using independent preparations of dyelabeled PG, but because of the different extents of dye-labeling, the data were not pooled.
(C) Purified His6-RlpA solubilizes dye-labeled PG sacculi from a ΔrlpA mutant.
Reaction conditions as above except that incubation was at 30°C for 2 hours (grey bars)
or 18 hours (black bars). The dashed line at 0.05 AU is provided to facilitate comparison
of the mutant proteins. (D) Identification of small PG fragments released by His6-RlpA.
Unlabeled PG sacculi from a ΔrlpA mutant were incubated with buffer (untreated) or
His6-RlpA (RlpA-treated). Reaction mixtures were centrifuged to separate residual
insoluble PG pellets from soluble PG fragments released into the supernatant. The pellet
and supernatant fractions were analyzed by reverse-phase HPLC. Peaks eluting from the
HPLC column are numbered as in Figure 3.8, except for two unique peaks labeled “a”
and “b” that were identified by tandem mass spectrometry (Table 3.6). Unreduced refers
to a sample that was not treated with borohydride. All chromatograms are graphed on the
same vertical (A206 nm) scale.
91
A
C
D
B
92
Figure 3.10. RlpA digests PG sacculi from a ΔrlpA mutant. Unlabeled PG sacculi
isolated from MJ1 (WT) or MJ24 (ΔrlpA) after growth in LB0N were incubated with
His6-RlpA. Reaction mixtures were centrifuged to separate residual insoluble PG from
soluble fragments released into the supernatant. Both fractions were subjected to
muropeptide analysis. Peaks are numbered and were characterized as described in the
legends to Figures 3.8 and 3.9.
93
94
Figure 3.11. RlpA does not cleave isolated tetrasaccharide. A portion of the P5 product
(NAG-NAM-NAG-NAMol) that had been isolated by RP-HPLC for mass spectrometry
analysis was divided into three aliquots and incubated with buffer (negative control), 4
μM His6-RlpA or 4 μM His6-RlpA(D157N) (another negative control). Reaction
mixtures were analyzed by RP-HPLC.
95
96
Figure 3.12. Amidase-treatment of PG renders it susceptible to subsequent cleavage by
His6-RlpA. Dye-labeled sacculi from a wild-type E. coli strain were incubated overnight
with buffer (as a control; filled squares “untreated”), 1 μM His6-AmiD (filled triangles)
or 1 μM His6-RlpA (open circles). These substrates were then incubated with 4 μM His6RlpA (A) or 4 μM His6-AmiD (B). A representative experiment is shown with one
replicate per time point. (C) Reproducibility of the assay. Dye-release was read after
480 min of incubation as indicated in (A) and (B). The values shown are the mean and
standard deviation of 4 separate experiments done with two preparations of dye-labeled
sacculi.
97
A
B
C
98
Figure 3.13. RlpA activity is potentiated by AmiD. This is a companion to Figure 3.12
and shows additional controls. Dye-labeled sacculi from a wild-type E. coli strain were
incubated overnight with buffer (“untreated”), 1 μM His6-AmiD or 1 μM His6-RlpA, as
indicated. These sacculi preparations were then incubated with 4 μM His6-RlpA, His6RlpA(D157N), His6-AmiD or buffer. Dye-release was read after 480 min of incubation.
99
100
Figure 3.14. Sequence analysis of RlpA. (A) Identification of conserved residues in the
RlpA-like DPBB domain targeted for mutagenesis. Mutagenized residues are bolded and
highlighted in grey. Conserved amino acids are indicated below the sequence with *
(invariant), : (highly conserved) and . (moderately conserved). Sequences were aligned
using Clustal Omega (Sievers et al., 2011) with default parameters. Sequences of RlpA
were obtained from: P. aeruginosa UCBPP-PA14 protein PA14_12090 residues 100-194,
Vibrio parahaemolyticus RIMD 2210633 protein VP0720 residues 84-178, Yersinia
pestis Z176003 protein YPZ3_2296 residues 65-171, Klebsiella pneumoniae 342 protein
KPK_3908 residues 79-170, E. coli K-12 MG1655 RlpA residues 79-171, and
Caulobacter crescentus ATCC 19089 protein CC_1825 residues 67-161. (B) Sequence
of the active site from RlpA and MltA aligned with other suspected lytic
transglycosylases. The catalytically important D157 of RlpA (this study) is bold and
highlighted in red. The catalytic D308 of MltA (van Straaten et al., 2007) is bold and
highlighted in blue. Conserved amino acids are indicated below the alignment as in (A).
To produce this alignment, we used PHYRE to model the DPBB domain of P.
aeruginosa RlpA onto the structures of PA4485, MltA and YoaJ. The relevant portions
of these sequences, together with the corresponding region from E. coli RlpA, were then
aligned using Clustal Omega (Sievers et al., 2011) with default parameters. The Clustal
alignment conformed to the PHYRE models. The sequences shown are: P. aeruginosa
UCBPP-PA14 protein PA14_12090 residues 152-174; E. coli K12 MG1655 RlpA
residues 131-153; P. aeruginosa PAO1 PAO1-UW protein PA4485 residues 84-106; E.
coli K12 MG1655 MltA residues 292-314 (numbering is for the mature protein, after
removal of the signal sequence); and B. subtilis 168 YoaJ residues 90-112.
101
A
B
P.
V.
Y.
K.
E.
C.
aeruginosa
parahaemolyticus
pestis
pneumoniae
coli
crescentus
MVGTASWYGTKFHGQATANGETYDLYGMTAAHKTLPLPSYVRVTNLDEKGRASWYGKKFQGHLTSNGEIYDMYSMTAAHKTLPLPSYVKVTNTDQIGLASSYGEEARGNTTATGEIFDPNALTAAHPTLPIPSYVRVTNVSQAGFAAIYDAEPNSNLTASGETFDPTQLTAAHPTLPIPSYARITNLAQAGLAAIYDAEPGSNLTASGEAFDPTQLTAAHPTLPIPSYARITNLAVVGIGSWYGEQFHNRKTSNGEIFDMNLPSAAHKTLPLPSLVEVTNLD* .: * :
. *:.** :*
:*** ***:** ..:**
P.
V.
Y.
K.
E.
C.
aeruginosa
parahaemolyticus
pestis
pneumoniae
coli
crescentus
NGKSVIVRVNDRGPFYSDRVIDLSFAAAKKLGYAETGTARVKVEGIDP
NGKTTVVRVNDRGPFHDGRIIDLSYAAAHKLDVIKTGTANVEIEVISV
NGRQIVVRVNDRGPYTPGRVIDLSRAAADRLNISN--NTKVKIDFINV
NGRMIVVRINDRGPYGNDRVISLSRASADRLNTSN--NTKVRIDPIIV
NGRMIVVRINDRGPYGNDRVISLSRAAADRLNTSN--NTKVRIDPIIV
NGRKMILRVNDRGPFVGDRIIDLSKAAADELGYRRQGVARVRVKYVGP
**: ::*:*****:
*:*.** *:*..*
.
..:
:--
RlpA_P. aeruginosa
RlpA_E. coli
PA4485_P. aeruginosa
MltA_E. coli
YoaJ_B. subtilis
IVRVNDRGPFYSDRVIDLSFAAA
VVRINDRGPYGNDRVISLSRAAA
VVRINDRGPFRRGRIIDVSRKAA
LMVALDVGGAIKGQHFDIYQGIG
TVYVTDLYPEGARGALDLSPNAF
:
*
:.:
102
Figure 3.15. Other PG hydrolases: SltB1, MltB1 and AmiB. (A) Division phenotypes of
lytic transglycosylase mutants do not mimic ΔrlpA. Cells grown at 37°C in LB or LB0N
to an OD600 ~0.5 were fixed, stained with the membrane dye FM4-64 and photographed
under fluorescence. The white bar represents 2 µm. Strains shown are MJ1 (WT), MJ24
(ΔrlpA), MJ34 (Δsltb1), MJ47 (Δmltb1), and MJ49 (Δsltb1Δmltb1). (B) Septal
localization of AmiB-mCherry does not require rlpA. Cells of MJ119
(WT/pJN105::amiB-mCherry) and MJ117 (ΔrlpA/ pJN105::amiB-mCherry) were grown
in LB0N to OD600 ~0.5 and photographed under phase (above) and fluorescence (below).
Filled triangles point to blebs where cells are lysing at division sites, perhaps provoked
by the AmiB-mCherry fusion. The fluorescence micrographs were inverted to better
visualize localization and blebbing.
103
A
B
(WT/pJN105::amiB-mCherry) (∆rlpA/pJN105::amiB-mCherry)
104
Figure 3.16. Structural comparison with MltA. RlpA threaded onto PA4485 (PDB
4AVR) was superimposed on MltA (PDB 2GAE) from E. coli (Moynie et al., 2013, van
Straaten et al., 2005). RlpA is colored in yellow and MltA in red. Despite having little
sequence similarity to one another, the overlays show the DPBB domains are very
similar. Aspartate residues shown to be important for catalysis in MltA (D297 and D308)
and the corresponding residues in RlpA (D157 and D168) are highlighted in blue and
magenta, respectively (van Straaten et al., 2005).
105
C
N
C
D297
D157
D308
N
D168
106
CHAPTER 4: IN VIVO AND IN VITRO STUDIES SUGGEST RLPA
OF ESCHERICHIA COLI IS NOT A LYTIC TRANSGLYCOSYLASE
Introduction
We demonstrated in chapter 3 that RlpA from P. aeruginosa (RlpAPa) is a lytic
transglycosylase that plays an important role in both cell division and cell shape. This
breakthrough prompted us to return to our studies of RlpA from E. coli (RlpAEc) to
determine if it also has lytic transglycosylase activity. On the surface, we would expect
the two RlpAs to have the same activity as they look and behave much the same; both
RlpAs (i) are found in a conserved locus containing genes involved in cell shape and PG
metabolism, (ii) have similar protein architectures [type II signal peptide, double-psi
beta-barrel domain (DPBB), SPOR domain] (Figure 4.1A), and (iii) localize to the
midcell and spots along the lateral wall (Gerding et al., 2009, Arends et al., 2010,
Jorgenson et al., 2014). A closer look at the sequence reveals, however, an important
difference. Specifically, a highly conserved aspartate in the DPBB domain of RlpAPa
(D168) that probably serves as the general acid/base during catalysis is a serine in RlpAEc
(S147) (van Straaten et al., 2005, Powell et al., 2006, van Straaten et al., 2007).
In this chapter we describe our efforts to determine if RlpAEc, like RlpAPa, has
lytic transglycosylase activity. We demonstrate that RlpAEc fails to rescue the P.
aeruginosa ∆rlpA phenotype despite localizing to sites of division. We go on to show
that changing D168 to serine in the DPBB domain of RlpAPa abolishes catalytic activity,
consistent with the postulated role of this residue in catalysis. However, efforts to “fix”
RlpAEc by changing S147 “back” to D were not successful—the S147D mutant protein
neither rescued division when produced in the P. aeruginosa ΔrlpA mutant nor
107
hydrolyzed PG sacculi when purified. Moreover, combining a ∆rlpAEc mutation with
deletions of other PG hydrolase genes in E. coli did not lead to synthetic phenotypes.
Taken together, our results suggest that E. coli RlpA is not a lytic transglycosylase. This
chapter also includes a further molecular characterization of RlpAEc, showing it is
trafficked to the OM and is present at about 600 molecules per cell.
Results
RlpAEc does not rescue a P. aeruginosa ∆rlpA mutant. We previously showed
that an rlpA deletion mutant of P. aeruginosa has morphological and viability defects
when grown in LB media lacking NaCl (hereafter referred to as LB0N) (Jorgenson et al.,
2014). Specifically, the ∆rlpA mutant formed chains of unseparated cells when grown in
LB0N broth and could not form colonies on LB0N plates. However, rlpA mutants of E.
coli do not have similar phenotypes (Gerding et al., 2009, Arends et al., 2010). A highthroughput screen of a large E. coli mutant library reported that a ΔrlpA mutant has
slightly increased sensitivity to a few antibacterial compounds such as carbenicillin and
SDS (Nichols et al., 2011), but we have not been able to reproduce these results (data not
shown).
To determine if RlpAEc has similar activity to RlpAPa, we first asked whether the
P. aeruginosa ∆rlpA mutant could be rescued by rlpAEc expressed from a plasmid. To
address this, we converted our rlpAPa-mCherry expression construct described previously
[pDSW1518 (Jorgenson et al., 2014)] to express rlpAEc-mCherry instead. In the
modified plasmid, the sequence corresponding to the mature protein of RlpAPa (residues
27-341) was replaced with the sequence to the mature protein of RlpAEc (residues 18362). The upstream sequence and the type II signal sequence from rlpAPa were retained
108
to allow for normal expression and processing. To test functionality, the rlpAEc-mCherry
expression plasmid was transformed into the P. aeruginosa ΔrlpA background. Although
RlpAEc-mCherry was produced in normal amounts and localized to sites of division
(Figure 4.1B and 4.2), it did not rescue the ∆rlpA mutant; cells harboring the expression
plasmid failed to form colonies on LB0N plates and looked chained in LB0N broth
(Figure 4.1C and 4.2).
RlpAEc does not exhibit PG hydrolase activity in vitro. We purified His6RlpAEc and tested its ability to degrade PG in solution using as substrate purified E. coli
sacculi labeled with the dye Remazol Brilliant Blue (RBB-PG) (Zhou et al., 1988).
However, incubation of the E. coli protein with RBB-PG did not result in dye release
(Figure 4.3A). As positive controls, we observed robust dye release using lysozyme and
weak dye release using purified His6-RlpAPa from P. aeruginosa (Figure 4.3A).
Pretreatment of the RBB-PG with an amidase to remove stem peptides stimulated dyerelease in the case of His6-RlpAPa, as shown previously (Jorgenson et al., 2014), but even
this substrate was refractory to digestion with the E. coli protein (Figure 4.3B).
Next, we explored an alternative assay format named zymography that has been
reported to reveal PG hydrolase activities in at least two cases where activity was not
observed using solution-based assays (Gutierrez et al., 2010, Bartual et al., 2014).
Briefly, 2 μg of purified proteins were run on two SDS-PAGE gels, one containing 0.5%
Micrococcus lysodeikticus cells and one without. After renaturation, the M. lysodeikticus
gel was photographed against a black background to visualize zones of clearing (PG
degradation) and then stained with methylene blue to better visualize clearing. Not
surprisingly, His6-RlpAEc did not show convincing clearing on zymography gels (Figure
109
4.4). Taken together, these results suggest that RlpAEc probably does not have PG
hydrolase activity.
Residue D168 in the DPBB is necessary for RlpAPa function in vivo. The
DPBB domain of RlpAPa has structural similarity to another lytic transglycosylase, MltA
of E. coli (Jorgenson et al., 2014). In MltA, there are two important aspartates in the
catalytic pocket, D297 and D308, the latter which acts as the general acid/base during
catalysis (van Straaten et al., 2005, Powell et al., 2006, van Straaten et al., 2007). These
residues align with D157 and D168 of RlpAPa. Though we previously showed that D157
is important for catalysis in RlpAPa (Jorgenson et al., 2014), we did not investigate D168.
We therefore constructed and characterized D168N and D168S mutants of RlpAPamCherry. Although both mutant proteins were produced at wild type levels (Figure
4.1B) and localized to sites of division (Figure 4.2), neither rescued the ∆rlpA mutant on
LB0N as assayed by plating and microscopy (Figure 4.1C and 4.2). These results are
consistent with the inference that D168 is the catalytic acid/based in RlpAPa.
A S147D substitution in the DPBB of RlpAEc is not sufficient to restore
catalytic activity. After demonstrating that D168 is necessary for RlpAPa activity, we
reasoned that the failure of RlpAEc to function as a PG hydrolase in vivo and in vitro
might be due to the fact that this residue is a serine in the E. coli protein. We therefore
constructed and characterized a S147D mutant of RlpAEc. However, RlpAEc(S147D)mCherry did not rescue the ΔrlpA phenotype in P. aeruginosa, even though it localized to
sites of division (Figure 4.1C and 4.2). Consistent with this result, the S147D protein
also failed to digest untreated or amidase-treated RBB-PG in dye-release assays (Figure
4.3A and B), and failed to digest M. lysodeikticus cells in a zymogram (Figure 4.4). It
110
must be noted that the DPBB domains from P. aeruginosa and E. coli exhibit about 50%
amino acid identity, so, in retrospect, it is perhaps not too surprising that a single change
of S147 is not sufficient to confer lytic transglycosylase activity on the E. coli protein.
An rlpAEc mutation does not exhibit synthetic phenotypes in combination
with PG hydrolase mutations in E. coli. As noted above, E. coli ΔrlpA mutants do not
exhibit any morphological defects (Gerding et al., 2009, Arends et al., 2010). Because E.
coli PG hydrolases are notorious for their redundancy (Heidrich et al., 2001, Heidrich et
al., 2002, Priyadarshini et al., 2007), we considered the possibility that the lack of
division or shape defects for E. coli ΔrlpA might reflect the ability of another PG
hydrolase(s) to substitute for RlpA. To explore this notion, P1-transduction was used to
construct a set of mutants lacking rlpA and various lytic transglycosylases or amidases.
These mutants were initially tested for growth defects when streaked onto LB plates, but
no such differences were observed (data not shown). To assay the mutants for subtle
morphological abnormalities, they were grown to mid-log phase in LB and examined by
fluorescence microscopy after staining the cells with the membrane dye FM4-64 to
facilitate visualization of cell contours and constrictions. Again, however, no phenotypic
changes attributable to ΔrlpA were observed (Tables 4.1-4.3). For example, consistent
with a previous report (Heidrich et al., 2002), an E. coli strain lacking three LT’s
(ΔmltCDE) had a subtle chaining phenotype, while a strain lacking five LT’s
(ΔmltACDEΔslt) had a somewhat more severe chaining phenotype. But deleting rlpA did
not exacerbate the division defect of either mutant. Similarly, as noted previously
(Heidrich et al., 2001, Priyadarshini et al., 2007), mutants lacking one amidase (ΔamiA or
ΔamiC) exhibited a subtle chaining defect, while a mutant lacking two amidases
111
(ΔamiAC) had a pronounced chaining defect. However, none of these defects were
accentuated upon introduction of an rlpA deletion.
RlpAEc is an OM lipoprotein that is present at 600 molecules per cell. During
the course of our studies, we decided to carry out a more detailed molecular
characterization of RlpAEc. We first investigated the membrane localization of RlpAEc.
We were motivated to do this since several online databases (e.g., Ecocyc, EcoProDB,
and Uniprot) annotate RlpAEc as an IM protein despite the fact that residues in its lipobox
motif (CTSDD) indicate RlpAEc is sorted to the OM. The serine at the +2 position
(underlined) of the mature protein predicts that RlpAEc is trafficked to the OM by the Lol
pathway (Seydel et al., 1999). To explore the membrane localization of RlpAEc, we
adapted an in vivo plasmolysis assay (Lewenza et al., 2006). Plasmolysis (hyperosmotic
shock) causes the IM to retract from the PG cell wall and can be seen as bays (phase light
regions) at the cell pole and along the lateral wall by phase-contrast microscopy (Figure
4.5). Conversely, the contour of the OM remains unchanged when cells are plasmolyzed.
To determine the membrane localization of RlpA, an rlpAEc-mCherry fusion was
expressed from a plasmid in an E. coli strain that also expresses a GFP fusion to an inner
membrane protein, FtsI (Rodriguez-Tebar et al., 1985, Bowler & Spratt, 1989, Weiss et
al., 1999). When cells were grown in LB, the green and red fluorescence were
essentially superimposable, as expected because our microscope is not able to resolve the
inner and outer membranes (Figure 4.5). In contrast, after cells were suspended in high
sucrose, GFP-FtsI was visualized along the edge of the plasmolysis bays, consistent with
its IM localization, while the contour outlined by RlpA-mCherry was similar to untreated
cells, demonstrating that RlpA is associated with the OM (Figure 4.5).
112
To estimate the amount of RlpAEc per cell, we used quantitative Western blotting
as previously described (Weiss et al., 1997). Briefly, RlpAEc from a known number of
cells was compared to a standard curve of purified His6-RlpAEc. We estimated there to
be 619 ± 237 molecules of RlpAEc per cell (mean ± standard deviation; n = 3 biological
replicates) in strain MG1655 when grown in LB medium at 37°C (Figure 4.6). This
number is in agreement with a recent estimate of approximately 900 molecules per cell
based on ribosome profiling (Li et al., 2014).
Discussion
We have shown that RlpA from E. coli is an OM protein present at about 600
molecules per cell. More importantly, our findings argue RlpAEc is not a lytic
transglycosylase, in contrast to its P. aeruginosa counterpart, which digests glycan
strands that lack stem peptides (Jorgenson et al., 2014). Our conclusion rests on three
independent lines of evidence. (i) Sequence analysis: the probable catalytic aspartate in
RlpAPa(D168) is a serine in RlpAEc(S147). This substitution is not a sequencing artifact
or mutation that arose during domestication of E. coli K-12 because the residue is a serine
in all sequenced E. coli strains, Salmonella enterica and Klebsiella pneumoniae. (ii)
Enzymatic assays: RlpAEc did not hydrolyze PG in any of the three assay formats tested,
all of which elicited at least some activity from RlpAPa. (iii) Mutant phenotypes:
Whereas a P. aeruginosa ΔrlpA mutant grew in LB0N as chains of short, fat cells, an E.
coli ΔrlpA mutant exhibited no such phenotypic changes, even when ΔrlpA was
combined with deletions of other PG hydrolase genes. Moreover, RlpAEc failed to rescue
the P. aeruginosa ΔrlpA mutant even though it localized to division sites.
113
This leaves us with several questions. One is, what fraction of the ~5000 genes
annotated as RlpA in the Pfam database as of June, 2014 (Finn et al., 2014) are bona fide
lytic transglycosylases? Studies of MltA and now RlpAPa have demonstrated that two
aspartates in the DPBB domain are critical for lytic transglycosylase activity (van
Straaten et al., 2005, Powell et al., 2006, van Straaten et al., 2007, Jorgenson et al.,
2014). In the Pfam seed alignment of 127 RlpA-like DPBBs, the first and second
aspartates are conserved in 69% and 87% of the sequences, respectively. These trends
are reflected in the Pfam HMM logo for RlpA-like DPBBs, which is derived from >5000
sequences (Figure 4.7). Because the second aspartate is probably the catalytic acid/base,
the fact that it is conserved in close to 90% of the sequences suggests 90% of the
annotated RlpA’s have lytic transglycosylase activity. However, if both aspartates are
required for enzymatic activity, only ~60% of the annotated RlpAs are LT’s (because at
least one of the critical aspartates is missing in 39% of the sequences in the seed
alignment).
A second outstanding question is, what is the function of RlpA in E. coli? This
remains a matter of conjecture. The architecture of the protein, with an N-terminal lipid
embedded in the OM and a C-terminal SPOR domain that can bind PG suggests RlpAEc
could help tether the OM to the PG sacculus. However, if this were the case, we would
expect a ΔrlpA mutant to exhibit OM blebbing and permeability defects that render these
mutants hypersensitive to toxic compounds like rifampicin and deoxycholate (Heidrich et
al., 2002, McBroom et al., 2006, Chimalakonda et al., 2011). We have not observed
either of these phenotypes, despite some effort to uncover them. Of note, we were not
able to reproduce a report that ΔrlpA renders E. coli slightly more sensitive to the
114
antibiotic carbenicillin (Nichols et al., 2011). Another potential function for RlpAEc is as
an assembly factor that helps to maintain the integrity of the multiprotein complex known
as the septal ring or divisome, although the lack of a division defect in an E. coli ΔrlpA
mutant argues against this.
It is interesting to note that RlpAEc is not the first septal ring protein that appears
to have started out in an evolutionary sense as an enzyme but has since lost that function
(Figure 4.8). At least three other periplasmic septal ring proteins fall into this category:
SufI, EnvC and NlpD. SufI (also called FtsP) is a member of the multicopper oxidase
family. Proteins in this family typically use copper atoms to oxidize small organic
molecules [reviewed in (Nakamura & Go, 2005)]. However, sequence alignments
revealed early on that SufI lacks key copper binding residues, suggesting it could not
function as a multicopper oxidase (Stanley et al., 2000). This inference was reinforced
when the crystal structure of SufI revealed that the catalytic pocket found in other
members of this enzyme family is occluded (Tarry et al., 2009). Although SufI is clearly
linked to cell division by its localization to the septal ring and several mutant phenotypes,
the specific function of SufI during cytokinesis remains obscure (Samaluru et al., 2007,
Tarry et al., 2009). EnvC and NlpD are members of the LytM family of
metalloendopeptidases (Pfam Peptidase_M23) [reviewed in (Firczuk & Bochtler, 2007)].
The best-characterized LytM proteins are PG hydrolases (e.g., lysostaphin and LytM
from Staphylococcus aureus) (Browder et al., 1965, Ramadurai et al., 1999). However,
the LytM domains of EnvC and NlpD lack key catalytic residues, and, at least in E. coli,
these proteins serve as allosteric activators of cell wall amidases (Uehara et al., 2010).
115
Table 4.1. Morphological parameters of lytic
transglycosylase mutants lacking rlpA in E. coli
% of cells w/indicated
number of constrictions
0
1
>1
WT
353
3.4 (0.8)
78
22
0
∆rlpA
375
3.5 (0.9)
79
21
0
∆mltCDE
419
4.4 (1.2)
50
50
0
∆rlpA ∆mltCDE
471
3.9 (1.2)
57
43
0
∆mltACDE ∆slt
278
5.8 (2.1)
29
64
7
∆rlpA mltACDE ∆slt
303
5.7 (2.0)
26
64
10
a
Strains shown are (in the order listed) EC251, EC3183, EC3745, EC3747, EC3702, and
EC3704.
Genotypea
b
No. of cells Avg Length,
evaluated
(SD)b
End-to-end length, regardless of the number of constrictions.
116
Table 4.2. Morphological parameters of single
amidase mutants in combination with ΔrlpA in E. coli
% of cells w/indicated
number of constrictions
0
1
>1
WT
369
3.3 (0.8)
85
15
0
∆rlpA
382
3.7 (1.1)
78
22
0
∆amiA
453
4.6 (1.2)
73
26
1
∆amiC
396
3.4 (0.9)
64
36
0
∆rlpA∆amiA
421
4.9 (2.1)
71
29
0
∆rlpA∆amiC
338
3.3 (1.2)
67
33
0
a
Strains shown are (in the order listed) EC251, EC3183, EC3433, EC3437, EC3439, and
EC3443.
Genotypea
b
No. of cells
evaluated
Avg Length,
(SD)b
End-to-end length, regardless of the number of constrictions.
117
Table 4.3. Morphological parameters of double
amidase mutants in combination with ΔrlpA in E. coli
% of cells w/indicated
number of constrictions
0
1
WT
333
2.2 (0.5)
98
2
∆rlpA
552
1.9 (0.4)
97
3
∆amiAC
335
4.3 (2.8)
36
35
∆rlpA∆amiAC
366
4.6 (3.2)
34
37
a
Strains shown are (in the order listed) EC251, EC3183, EC3486, and EC3492.
Genotypea
b
No. of cells
evaluated
Avg Length,
(SD)b
>1
0
0
29
29
End-to-end length, regardless of the number of constrictions. The cells are shorter
because these values are from overnight cultures.
118
Figure 4.1. Phenotypes of ΔrlpA expressing rlpAEc or mutants of rlpA with lesions in the
DPBB domain. (A) Cartoon depicting the domain architecture of RlpA from E. coli K12 and P. aeruginosa UCBPP-PA14. S, type II signal sequence (including the lipobox
sequence); DPBB, RlpA-like double-psi beta-barrel domain; SPOR, SPOR domain; these
domains are connected by linker regions depicted as thin lines. Numbers below the
diagrams refer to amino acids identified as domain boundaries as annotated in the Pfam
database (Finn et al., 2014). The two RlpAs share 46% sequence identity in the DPBB
domain, 25% sequence identity in the SPOR domain, and 26% sequence identity overall.
There is no significant homology between the linker regions. The position of the lipid
modified cysteine is noted above each graphical view. (B) Western blot with antimCherry sera. Molecular weight markers are indicated to the left of the blot. The
predicted masses are 61 kDa for RlpAPa-mCherry proteins and 63 kDa for RlpAEcmCherry proteins, assuming removal of the signal sequence. Note that RlpAEc has
previously been shown to migrate slower than its predicted molecular mass (Takase et
al., 1987). (C) Plating efficiency. Tenfold serial dilutions of cells with the indicated
genotypes were spotted on LB Gent (left) and LB0N Gent (right). Plates were incubated
overnight at 37°C and then photographed. Strains shown are ∆rlpA/pJN105 (MJ26),
∆rlpA/pDSW1518 expressing RlpAPa-mCherry (MJ40), ∆rlpA/pDSW1545 expressing
RlpAPa(D157N) -mCherry (MJ71), ∆rlpA/pDSW1554 expressing RlpAEc-mCherry
(MJ73), ∆rlpA/pDSW1676 expressing RlpAPa(D168N) -mCherry (MJ131),
∆rlpA/pDSW1694 expressing RlpAPa(D168S) -mCherry (MJ133), and ∆rlpA/pDSW1695
expressing RlpAEc(S147D)-mCherry (MJ138).
119
A
C18
E. coli
S
1
DPBB
20
362
SPOR
78
167
287
360
C27
341
P. aeruginosa
1
29
99
189
262
339
B
Dilution factor
C
-1
∆rlpA/pJN105::rlpAPa-mCherry
∆rlpA/pJN105
∆rlpA/pJN105::rlpAPa(D157N)-mCherry
∆rlpA/pJN105::rlpAPa(D168N)-mCherry
∆rlpA/pJN105::rlpAPa(D168S)-mCherry
∆rlpA/pJN105::rlpAEc-mCherry
∆rlpA/pJN105::rlpAEc(S147D)-mCherry
-2
-3
-4
-5
-6
-1
-2
-3
-4
-5
-6
120
Figure 4.2. Function and localization of RlpA proteins. Strains grown in LB0N Gent at
37°C to an OD600 ~0.5 were imaged by phase contrast (left) and fluorescence (right)
microscopy. Arrows in (B) point to septal localization of RlpAPa-mCherry. The black bar
represents 2 μm. ΔrlpA strains containing pJN105 derivatives shown are listed in the
legend to Figure 4.1.
121
122
Figure 4.3. RlpAEc does not have PG hydrolase activity in solution. Dye-labeled sacculi
from a wild-type E. coli strain were incubated overnight with buffer or 1 μM His6-AmiD.
These substrates were then incubated with the indicated proteins at 4 µM. Dye-release
from untreated (A) and amidase treated (B) sacculi was assessed after overnight
incubation (>18 hours) at 37°C. Lysozyme (LZ) served as the positive control. P.a._WT
= His6-RlpAPa. P.a._D157N = His6-RlpAPa(D157N). E.c._WT = His6-RlpAEc.
E.c._S147D = His6-RlpAEc(S147D). The values shown are the mean and standard
deviation of a representative experiment done in triplicate.
123
A595nm
A
B
0.35
0.30
0.25
0.20
0.15
0.10
0.05
0.00
0.50
A595nm
0.40
0.30
0.20
0.10
0.00
124
Figure 4.4. RlpAEc does not have activity in a renaturing gel electrophoresis
(zymography) assay. 2 μg of the indicated proteins were run on two SDS-PAGE gels.
Top panel: Gel without PG was stained with Coomassie blue to visualize migration and
purity of proteins. Middle panel: Gel containing M. lysodeikticus cells was
photographed against a black background after incubating in renaturing solution
overnight. Bottom panel: Same gel after staining with methylene blue to better visualize
zones of clearing. Lysozyme (LZ) served as a positive control. Bovine serum albumin
(BSA) served as a negative control. P.a._WT = His6-RlpAPa. P.a._D157N = His6RlpAPa(D157N). E.c._WT = His6-RlpAEc. E.c._S147D = His6-RlpAEc(S147D). The
predicted molecular masses are 14.3 kDa for LZ, 66.5 kDa for BSA, 35.1 kDa for His6RlpAPa proteins, and 36.7 kDa for RlpAEc proteins.
125
126
Figure 4.5. RlpAEc is trafficked to the OM. Cells grown in LB with 100 µM IPTG were
recovered and then resuspended in either LB (top panel) or plasmolyzed in 15% sucrose
(bottom panel). Cells were then spotted onto 1% agarose pads. To maintain plasmolysis,
cells in sucrose were spotted onto 1% agarose pads containing 15% sucrose. Arrows in
the bottom panel point to plasmolysis bays. Strain shown is EC3129.
127
128
Figure 4.6. Quantitative Western blot showing there are approximately 600 molecules of
RlpAEc per cell. (A) The amount of RlpAEc from 1.5 x 108 cells from strain EC251 (wild
type) was compared to a standard curve ranging from 0 ng to 4 ng of purified His6RlpAEc added to a ∆rlpAEc (EC3183) extract. (B) Quantification of chemiluminescence
from RlpAEc (closed squares) and His6-RlpAEc (open circles). The predicted molecular
mass of native RlpAEc (assuming removal of the signal sequence) and His6-RlpAEc is
35.7 kDa and 36.7 kDa, respectively. EC251 was loaded twice for reproducibility.
129
A
B
14000
12000
AU
10000
8000
6000
4000
2000
0
0
1
2
3
RlpA, ng
4
5
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Figure 4.7. Pfam Hidden Markov model (HMM) logo of the RlpA-like DPBB domain.
D157 in RlpAPa and D137 in RlpAEc correspond to position 46 in the HMM logo, which
can either be an aspartate or an asparagine. D168 in RlpAPa and S147 in RlpAEc
correspond to position 57 in the HMM logo, which is primarily an aspartate. The HMM
logo is based on 5162 sequences (Finn et al., 2014) and was retrieved on June 4, 2014.
131
132
Figure 4.8. Conservation of the putative catalytic aspartate in RlpA from different
proteobacteria. (A) The catalytic aspartate is conserved in all lineages except the
gammaproteobacteria, where some organisms have D and others have S. (B) The
catalytic aspartate is a serine in E. coli and closely-related members of the family
Enterobacteriaceae. The * indicates the phylogenetic branch point for the Asp to Ser
substitution. The pink highlight unites organisms where this residue is a serine. Note
that in both (A) and (B) the branch lengths are arbitrary. The phylogenetic trees were
modeled after (Emerson et al., 2007) and (Baumler et al., 2013) and are based on
multiple criteria, including sequences of rRNA genes and conserved proteins. RlpA
sequences were retrieved for multiple representatives of each group, alignments were
constructed to identify the residue (Asp or Ser) that aligned with D168 of RlpA from P.
aeruginosa, and that information was transferred to the phylogenetic trees to generate the
figures. The criteria used to classify proteins as RlpA included the presence of a SPOR
domain and, in the case of the gammaproteobacteria, synteny. Outside of the
gammaproteobacteria, synteny breaks down. Note that many proteins annotated as RlpA
in the Pfam database share only the DPBB domain with RlpA of P. aeruginosa and E.
coli; these proteins were not included in this analysis because we were not sure if they
were legitimate orthologs.
133
A
Deltaproteobacteria (D)
Epsilonproteobacteria (D)
Alphaproteobacteria (D)
Gammaproteobacteria (D/S)
Betaproteobacteria (D)
B
134
CHAPTER 5: FUTURE DIRECTIONS
Our studies of RlpAPa have led to a deeper understanding of cell division and
regulation of PG hydrolases. These insights have raised many new questions. Most
notably, we are still left with the conundrum of what RlpA is doing in E. coli. Below, we
discuss several open questions and approaches to answer them.
E. coli
What sequence changes are necessary to restore PG hydrolase activity to
RlpAEc? We hypothesized that RlpAEc lacks lytic transglycosylase activity, in part,
because a critical aspartate (D168) in RlpAPa is substituted for a serine (S147) in RlpAEc;
however, simply changing this residue back to an aspartate in the E. coli protein failed to
restore catalytic activity. Therefore, other residues must be important for catalysis, but
which ones? The DPBB domains from E. coli and P. aeruginosa are about 50%
identical, so there are many possibilities. Obtaining and characterizing RlpAEc mutants
that have acquired lytic transglycosylase activity is a priority because this will provide
insights into the mechanisms of binding and/or catalysis. A promising approach would
be to randomly mutagenize rlpAEc on a plasmid, transform into the P. aeruginosa ΔrlpA
strain, and select for rare survivors on LB0N. If mutants can be obtained, we would use
DNA sequencing to identify the changes and then purify the mutant proteins and
characterize them in vitro to verify that they now have lytic transglycosylase activity. An
especially interesting question will be whether the mutant proteins exhibit the preference
of RlpAPa for naked glycans. Because most lytic transglycosylases cut glycans that
contain peptide sidechains, it is possible that (some) mutant RlpAEc proteins may also.
135
This line of investigation could provide insights into which residues are important for
catalytic activity and substrate specificity. Similarly, one could mutagenize the RlpA
protein from P. aeruginosa and screen for variants that lyse E. coli when overproduced.
(We already know that overproduction of wild type RlpAPa does not lyse E. coli,
presumably because the activity of that protein is restricted to naked glycans.)
What is the function of RlpAEc? The central question regarding RlpAEc
remains, what is its role in the cell? Because RlpA is anchored in the OM by a lipid and
binds the PG via a SPOR domain, it seems well-suited to help anchor the OM to the PG.
This function might be especially important during constriction, which fits well with the
enrichment of RlpA in septal regions. Initial studies suggest an rlpA mutant does not
have obvious OM anchoring phenotypes such as increased sensitivity to compounds (e.g.,
rifampin and deoxycholate) that would indicate a disrupted OM barrier (data not shown).
However, two reports noted that an E. coli rlpA mutant might have subtle OM defects
(Nichols et al., 2011, Paradis-Bleau et al., 2014). Because E. coli is known to have
several proteins that contribute to OM anchoring, one potential explanation for the lack of
a strong rlpA phenotype is functional redundancy. To explore this hypothesis, I would
combine rlpA with deletions of genes that code for OM proteins that help with
anchoring—Lpp, NlpI, OmpA and Pal (Fung et al., 1978, McBroom et al., 2006, Leduc
et al., 1992, Park et al., 2012, Cascales et al., 2002, Parsons et al., 2006, Gerding et al.,
2007). The mutants would be examined by microscopy and for sensitivity to compounds
that ordinarily do not cross the OM very well such as bacitracin, deoxycholate, SDS, etc.
Besides having potential to identify a convincing function for RlpAEc, this line of
136
investigation could shed light on the important question of how invagination of the layers
of the cell envelope is synchronized.
P. aeruginosa
What is the function of PA4485? During the course of our studies, we became
aware of a second RlpA-like protein in P. aeruginosa, PA4485. RlpAPa and PA4485 are
similar in that they both contain a type II signal sequence, are predicted to be trafficked to
the OM, and have an RlpA-like DPBB domain. Unlike RlpAPa, PA4485 does not contain
a C-terminal SPOR domain. Interestingly, there are two independent transposon mutant
libraries of P. aeruginosa, but neither set contains a mutant of PA4485 (Jacobs et al.,
2003, Liberati et al., 2006). On the surface, this would suggest PA4485 is essential in P.
aeruginosa. However, this would be very unusual as we are aware of only one instance
of an essential PG hydrolase, PcsB of S. pneumoniae (Sham et al., 2011). Thus, it would
be interesting to try to knock-out PA4485, determine whether expressing PA4485 can
rescue the P. aeruginosa ∆rlpA mutant, and to purify PA4485 and test it for PG hydrolase
activity in vitro.
How do the DPBB domain and the SPOR domain of RlpAPa work together?
We showed that RlpAPa has a strong preference for naked glycans (Jorgenson et al.,
2014), which are also the suspected binding site for the SPOR domain (Ursinus et al.,
2004, Gerding et al., 2009). This would suggest to us that the SPOR domain enhances
the enzymatic activity of RlpAPa. To test this, we would construct a SPOR-less
derivative of RlpAPa and determine its efficiency at digesting PG as compared to the fulllength protein. In addition, we would ask whether deleting the SPOR domain from
mutant RlpA proteins with lesions in the DPBB (E120A, D123A, and H131A) renders
137
these proteins no longer able to rescue division in vivo. An alternative approach would
be to swap the SPOR domain for a PG binding domain with a different specificity (e.g.,
LysM domain [Pfam 01476]) and compare the swap construct to the wild type in vitro
and in vivo (Buist et al., 2008, Visweswaran et al., 2011). Finally, having a crystal
structure of RlpAPa would provide information as to whether these two domains interact
with one another, and if so, give us a more detailed model for the function of RlpAPa.
Does RlpAPa work together with a specific amidase? A major question in the
field of PG metabolism is how the many PG hydrolases work together. Our studies
indicate that amidases and RlpAPa work together in an ordered and sequential fashion—
first amidases remove stem peptides, then RlpAPa degrades the glycan chains. What
remains unresolved is whether RlpAPa works in tandem with a specific amidase. There
are two candidates in P. aeruginosa—AmiA and AmiB. To explore the relationship of
these amidases to RlpAPa in the cell, we would test for protein-protein interactions and
synthetic phenotypes.
What is the enzymatic specificity of RlpAPa? Because RlpAPa represents an
unusual type of lytic transglycosylase, its preference for naked glycans warrants deeper
characterization. The availability of naked glycans (Harz et al., 1990) puts us in a
position to address two issues: what is RlpAPa’s glycan length preference and where does
it cut? We already know that RlpAPa does not cut tetrasaccharides (Jorgenson et al.,
2014), which suggests it requires a larger glycan, but how large? The fact that we saw
release of tetra- and hexasaccharides means RlpAPa is an endo enzyme (Jorgenson et al.,
2014), but we did not follow the distribution of cuts. Also, because we do not know the
limits of detection of our assays, we do not know if RlpAPa exclusively generates 1,6-
138
anhydro NAM ends or whether it also produces a small amount of NAM, similar to
lysozyme. Using size exclusion chromatography on naked glycans (generated by
digesting with amidase to completion), together with mass spectrometry should resolve
these issues.
Is RlpAPa important for virulence? Cell shape is an important determinant of
the virulence of several pathogens [reviewed in (Young, 2006, Wyckoff et al., 2012,
Frirdich & Gaynor, 2013)]. The cell shape defects we noted for an rlpA mutant in P.
aeruginosa suggest rlpA might be important for virulence in this pathogen. Several
models have been described for P. aeruginosa infection and could be used to test our
mutants of rlpA in organisms ranging from fruit flies, to zebrafish, to mice (D'Argenio et
al., 2001, Clatworthy et al., 2009, Hoffmann et al., 2005). If rlpA is important for
virulence, it might be a useful drug target.
Why salt? A longstanding question in the field of cell division centers on the
observation that many cell division phenotypes are salt (osmotic) dependent (Ricard &
Hirota, 1973, Storts et al., 1989, Schmidt et al., 2004, Addinall et al., 2005, Reddy, 2007,
Samaluru et al., 2007, Karimova et al., 2012, Ransom et al., 2014). Case-in-point, an
rlpA mutant looks normal in LB containing salt but grows as chains of cells that
eventually lyse when the salt is removed from the growth media (Jorgenson et al., 2014).
However, it is not fully understood why these phenotypes are dependent on the presence
of salt. Studies have shown that the cell counteracts the loss of salt in its environment
(increase of turgor) by decreasing the intracellular pool of compatible solutes (e.g.,
glycine betaine) in its cytoplasm [reviewed in (Kempf & Bremer, 1998)]. The resulting
change in chemical composition of the cell may have profound effects on protein
139
stability, protein-protein interactions and/or enzymatic activities. However, it is not clear
whether osmotic balance pertains in the case of cell division mutants because many saltremedial mutants have defects in periplasmic proteins. More studies are needed to
investigate the basis of these salt dependent phenotypes because their ubiquity suggests
an important aspect of cellular physiology that remains to be understood.
140
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