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FINAL APPROVAL OF DISSERTATION Doctor of Philosophy in Biomedical Sciences
Translational Control in Escherichia coli: Hfq and PvuII
Submitted by: Meenakshi Kaul Kaw
In partial fulfillment of the requirements for the degree of Doctor of Philosophy in Biomedical Sciences
Examination Committee
Major Advisor: Robert Blumenthal, Ph.D.
Academic Randall Worth, Ph.D. Advisory Committee: Sonia Najjar, Ph.D.
Eric Lafontaine, Ph.D.
Kevin Pan, Ph.D.
Senior Associate Dean College of Graduate Studies Michael S. Bisesi, Ph.D.
Date of Defense: April 18, 2007
Translational Control in Escherichia coli: Hfq and PvuII
Doctor of Philosophy
Meenakshi Kaul Kaw
The University of Toledo College of Medicine
2007 DEDICATION
To my parents, Dr. H. K. and Susheela Kaul, your prayers, love and constant encouragement has given me strength to turn this hardest personal challenge into reality.
To my husband, Dr. Dinkar Kaw, for your love and unwavering support which has always been the most motivating forces in my life.
ii ACKNOWLEDGEMENTS
I am grateful for my major advisor, Dr. Robert Blumenthal for giving me his support, and opportunity of working in his laboratory. His close guidance, freedom, trust and respect over the years has been priceless. I also thank you for you for the scientific advice, as well as proffesional and personal advice in tough times.
I would also like thank my previous advisor, Dr. Darren Sledjeski for his support, time, patience and scientific experties that helped to attain my goals.
I would aslo like to thank my committee members for their invaluable time and patience and scientific advice they shared from time to time: Dr. Sonia
Najjar, Dr. Eric Lafontaine, Dr. Randal Worth and my unofficial member Dr. Mark
Wooten.
I thank the State of Ohio and The University of Toledo for the pre-doctoral fellowship and the tution support provided to me for four and a half years of my study.
I also thank the NIH for the Hfq project support. The material for PvuII project is based upon the work supported by NSF under Grant No. 0516692 and under the guidance of Dr. R.M. Blumenthal.
iii I would also like to thank all the laboratory and Club Coli members, past and present, who have contributed to helpful scientific discussions. To name just few, Robert Lintner, John Lazarus, Iwona Mruk and Pankaj Mishra.
Additionally I would like to thank the members of Department of Medical
Microbiology and Immunology, especially Joyce Rodebaugh, Sharon Ellard, Sue
Payne and Diane Ammons.
I would like to thank my beautiful daughter Meesha Kaw for her immense support and patience, my mother inlaw Dr. Durga Kaw, my sisters Er. Kamakshi
Kaul and Dr. Vinay Kaul.
I would also like to thank my nephews Akash and Smish, nieces Arshee and Anapurna. Thank you for your love for me.
Last but not least I would like to thank my cousin Er. Savita Bhan Wakhloo for her constant love and support.
iv TABLE OF CONTENTS
Page
Dedication …………………………………...... ii
Acknowledgements …………………………………………………. iii
Table of contents …………………………………………………. v
Introduction …………………………………………………. 1
Literature …………………………………………………. 5
Manuscript I …………………………………………………. 40
Manuscript II ………………………………………………….. 90
Discussion ………………………………………………….. 143
Bibliography ………………………………………………….. 149
Abstract ………………………………………………….. 186
v INTRODUCTION
In prokaryotic world, the majority of gene regulation appears to occur at
the level of transcript initiation (Boelens and Gualerzi, 2002), in part because
transcription and translation are concurrent. Neverthless the fastest way for the
repertoire of cell proteins adapt to change in the environment or the cellular
states is by regulating the translation. Translation can be repressed or enhanced
by displacement, entrapment or competition (Schlax et al., 2001). During
conditions of stress in the form of changes in pH, temperature, nutrient availbility,
or osmolarity, cells often go into stationary phase (Loewen et al., 1998; Hengge-
Aromis 1996). Various global regulators control the gene expression by different
mechanisms that help the cell adapt to the unfavorable environment, and escape
death.
Hfq protein from Escherichia coli is a global regulator central to multiple
regulatory processes (Muffler, et al., 1997; Tsui, et al., 1994). It is a key player in
post-transcriptional regulation that acts by facilitating base-pairing of bacterial
noncoding RNAs (ncRNAs) to their mRNA targets sites. (Schuppli, et al., 1997;
Muffler, et al., 1997; Tsui, et al., 1997; Wassarman, et al., 2001). Evidence suggests that Hfq acts as a chaperone and modulates RNA structure (Moll, et al.,
2003). Studies by Lease and Woodson, (2004) and E. Espinosa, P. Mikulecky and A. Feig, (personal communication) demonstrated that Hfq binds DsrA and rpoS RNAs independently and form stable ternary complexes. DsrA is a small 85
1 nucleotide ncRNA that is produced in response to low temperature and regulates
translation of two global regulatory proteins, RpoS and HNS (Sledjeski and
Gottesman, 1995) while rpoS mRNA specifies an alternative RNA polymerase sigma factor that plays a major role in transcribing stationary phase-associated genes (Pratt and Silhavy, 1998; Hengge-Aronis, 1996; Bougdour et al., 2004).
The existing data imply that more than one site is required for Hfq to promote base-pairing between ncRNAs and their targets. We addressed these questions by mutational analysis of Hfq. My experiments in this study were designed for in vivo mutationally altered Hfq. I used reporter assays and RNA life time experiments. Combining these results with in vitro analyses by our collaborators at Indiana University, we hypothesized that there are at least two independent sites on the Hfq hexamer, and close proximity of these RNA-bound sites facilitate base-paring between the bound RNAs.
In addition to the role of a global regulator such as Hfq, translational control can occur via cis-acting gene specific signals. I have studied this in genes for which regulation is particularly critical. Restriction Modification (RM) systems have a wide distribution in bacteria and archaea. RM systems include two components, a restriction endonuclease (REase) and a methyltransferase
(MTase). I studied RM systems in which the REase and MTase are independent proteins (Hedgpeth, et al., 1972; Schildkraut I et al., 1984 Roberts, et al., 2003).
2 A subset of the type II RM systems include a third protein designated as
controller or C protein that plays a regulatory role. PvuII is a C- dependent type II
RM system derived from the Gram negative rod Proteus vulgaris (Gingeras et al,
1981) that was cloned and can be studied in E. coli (Blumenthal, et al., 1985).
When RM systems move into a new host cell, it is crucial that the new host’s
DNA be methylated protectively before REase expression occurs. The C
proteins, such as C.PvuII play a central role in mediating this delay, and pre-
expressing C.PvuII in a cell prevents transformation by the intact RM system
(presumably due to premature REase production; (Vijesurier et al., 2000)
Previous studies from our laboratory (Tao et al 1991; Tao et al., 1992; Vijesurier et al., 2000; Knowle et al., 2005) demonstrated the transcriptional regulation of
REase by the C.PvuII.
However, work from other laboratories also indicated that translational control by C protein can occur. A study of eco72IC demonstrated that C protein translationally controls expression and act as an activator of REase gene expression (Rimiseliene, et al., 1995) while C protein from the Esp13961 also demonstrated that it activates its own expression and its downstream REase gene by mechanism of translational coupling (Cesnaviciene, et al., 2003).
Work from our laboratory indicated that C protein regulates REase expression via a transcriptional mechanism, but it is not clear whether this is sufficient to protect the cell. We investigated if there is some type of additional
3 translational-level control. Since there is an overlap region in this subset of C- controlled RM systems, we investigated whether this implied to some kind of translation control. In the PvuII RM system, there are two characteristics suggestive of translation regulation. First in PvuII and most C-dependent RM systems there is an overlap or close proximity between the C gene and the downstream R gene. Second in PvuII RM system this overlap is preceded by potential hairpins, one of which would occlude the ribosomal binding site (RBS)
The set of experiments designed to analyze this study included cloning, site- directed mutagenesis, and beta-galactosidase assays.
4 LITERATURE REVIEW
Part I: Translational control of genes in E. coli.
Part II: Hfq as a regulator of translation mediated by DsrA RNA in E. coli
Part III: C.PvuII as a regulator of translation of PvuII RM system in E. coli
5 PART I
Brief history of translation
The history of the evolution of the translationary machinery seems to have
started with resolving the mysteries behind the evolutionary transition from RNA to DNA. The “RNA world” (Gilbert 1986) hypothesized that RNA was the major biological catalyst and genetic material before protein and DNA (Woes 1967;
Crick 1968; Orgel 1968). The prediction from the RNA world model was that in the RNA world, protein synthesis evolved from RNA (Poole et al., 1998; Noller,
2004) before the three domains-namely archaea, bacteria, and eukaryotes separated out (Poole and Logan 2005). Futher research findings supported the evolution of protein from RNA and are accepted world wide. It is assumed that during the evolution of enzymatic activity, catalysis is transferred from RNA to ribonucleoprotein (RNP) to protein, the first proteins in the "breakthrough organism" (the first to have encoded protein synthesis) would be nonspecific chaperone-like proteins rather than catalytic that bind and stabilize the RNA structures (Poole et al., 1998; Noller, 2004). This still did not answer the question completely and researchers came with the possibility that translational apparatus evolved to expand the structural and the functional, capabilities of the RNA. This was supported by the rearrangements that occur in the three dimensional structure of RNA by binding of proteins, or peptides or other low molecular weight ligands affecting the overall transcriptional or translational regulation of mRNA
(Noller 2004). The origin of ribosomes occurs from preribosomal particles which
6 have only RNA, and the recruitment of ribosomal proteins occurs later in the event to enhance ribosomal assembly, speed and accuracy of translation, and mediate regulation of translation (Hoang et al., 2004).
Mechanism of translation initiation
In the translation process, the codon sequence on mRNA directs polypeptide chain synthesis. Initiation of translation in E. coli is mediated by three initiation protein factors IF1, IF2 and IF3. The process begins when the ribosomal, subunits 30S and 50S join to form 70S ribosome. Each subunit has three binding sites for tRNA, A (aminoacyl) that accept the incoming aminoacetylated tRNA, P (peptidyl) which bind the tRNA with nascent peptide chain, and the E (exit) site which holds the deacylated tRNA before it leaves the ribosome (Ramakrishnan, 2002).
The 30S subunit binds mRNA codon loops and anticodon stemloops of tRNA. It plays a significant role by monitoring the base-pairing between the codon on mRNA and the anticodon on tRNA during decoding of mRNA. The 50S catalyzes the binding of incoming amino acid on A-site tRNA to the growing peptide chain on the P-site tRNA (Ramakrishnan, 2002).
Process of initiation of translation
Initiation of translation is a central event in the cell that determines the efficiency of gene expression. In prokaryotes translation initiation occurs in
7 several steps. It begins with selecting of the mRNA translation initiation region
Met (mRNA TIR) as well as the initiator tRNA (fMet-tRNA f ) by 30S ribosomal subunit to form a ternary complex (Gulerzi and Pon, 1990). The 30S association complex either dissociates into its individual components or reforms the 70S by combining with the 50S. IF1 and IF3 fall off after the association complex is formed and 70S initiation complex starts to form. This stimulates intrinsic
Met GTPase activity of IF2, and placing fMet-tRNA f on the P-site, now IF2
dissociates itself from the complex. This enables the A-site on the 70S to be
freed to bind another elongator aminoacyl tRNA encoded by next mRNA codon.
Crystal structure analysis of the 30S subunit reveals that IF1 binding on its
A-site leads to extensive conformational changes in the 30S subunit (Carter et
al., 2001). A further study reveals two sites on IF2, IF2α and IF2β. The first site
aids in its binding to the 30S subunit and the second helps in the decoding step
and requires IF1 and GTP (Caserta et al., 2006). Similarly IF3 possesses two
domains (IF3C and IF3N) which are separated by a linker. IF3 uses its C domain
to bind to 30S subunit platform and uses N domain to bind to the P-decoding
region with its N domain. Like IF2, IF3 participates in translation fidelity, and
shifts equilibrium towards dissociation, so that 30S subunit is available for
translation initiation (Fabbretti et al., 2006). EF-Tu is the carrier of the ternary
complex, next an initiation dipeptide is formed, and subsequent to translocation
event (facilitated by EF-G) ribosome enters the next round of elongation phase of
translation (Boelens and Gualerzi, 2002; Selmer et al., 2006).
8 Start codon significance
The AUG start codon is reported to be the most efficient start codon.
When AUG was replaced by GUG or UUG, translation decreased by ~8-fold
(Sussman et al., 1996). AUU, a natural start codon for two E. coli genes is not as
efficient as UUG (Sussman et al., 1996). The molecular mechanism behind this
Met ternary complex formation of tRNA, ribosome and fMet-tRNA f is base pairing of the Shine Dalgarno (SD) sequence on mRNA with the anti SD sequence residing at 3’ end of 16S rRNA (Jacob et al., 1987; Steitz and Jakes 1975). In E. coli the majority of the genes analyzed so far show SD sequence with the exception of few (Shultzaberger et al., 2001) and is mostly present 5-8 nucleotides upstream of initiation codon. To be effective SD sequence does not need to be more than 5-6 base pairs (Munson et al., 1984).
The ribosomal binding site (RBS) ~15 nucleotides (nt) is present on either side of AUG (Steitz and Jakes, 1975) and extends 5’ of the SD sequence. It is proposed that the AU rich sequences prevent the formation of secondary structure, and enhance translation (Chen et al.,1994; Qing et al., 2003) and may explain the A rich sequence of E. coli mRNA at position -1 to -6 (Schauder and
McCarthy, 1989). Then begins the process of termination by presenting a stop codon to mRNA on the A-site, recognized by release factors (RF), UAA is recognized by both RF1 and RF2, however RF1 is for UAG and RF2 recognizes
UGA. The whole complex now initiates hydrolysis and peptide chain release from tRNA in the P-site. Ribosomal reassembly is brought about by ribosome
9 recycling factor (RRF) and requires EF-G (Janosi et al 1996). In addition the
expression of genes in E. coli also occurs via polycistronic transcripts, and the
translation of the downstream gene can be coupled to the upstream gene. In
most cases of coupled translation, the RBS from the downstream gene remains
obscured temporarily and may also form secondary structures. Sometimes
translation is permitted even if the downstream gene is leaderless. Translation
efficiency increases if the downstream gene possesses its own SD (Rex et al.,
1994).
Mechanism of Reinitiation by coupled translation
Reinintiation of translation starts once the ribosomes from the upstream gene are delivered to the following cistron, and within the two, the terminator codon of preceding gene overlaps with initiator codon of the next. This proximity facilitates reinitiation of translation e.g.UGAAUG (Sprengel etal 1985). In some of
these polycistronic mRNAs, as in most cases of coupled translation, the
downstream cistron has a RBS that is usable but is temporarily obscured by
secondary structures (Chen and Yanofsky, 2004).
Initially it was proposed that leaderless mRNA binds more stably to 70S
ribosome than to 30S subunits (O’Donnell and Janssen, 2002). Chemical cross-
linking studies demonstrated that in leaderless mRNAs translation begins in the
70S ribosome (Moll et al 2004). The 30S was required for initiation. IF3 is an
inhibitor of leaderless translation because it promotes dissociation of the 70S;
10 Met however IF2 promotes translation by stabilizing the binding of fMet-tRNA f
(Grill et al., 2000, Moll et al., 2002).
Regulation of translation by displacement, competition and entrapment
In prokaryotes or eukaryotes, the quickest way that the cell adapts to any environmental change or stress, is by regulating translation. Translation can be activated or repressed by rearrangement of the base-paired mRNA and its target gene, a step that will either obscure or open up the RBS, and determines the efficiency of translation. This rearrangement can be augmented or inhibited by the presence of other factors like mRNA binding proteins, small transacting
RNAs or the movement of the ribosome itself. Repression of translation via mRNA binding protein occurs when the repressor protein competes with the ribosomes to bind to RBS or close enough to occlude ribosomal entry (Jenner et al., 2005). Threonyl-tRNA synthetase binds to the operator site of its own mRNA, upstream of initiation codon, inhibiting ribosomal attachment and thus translation initiation. Base-pairing between the RBS and the repressor, inhibits binding of ribosome aggravating repression of the gene thrS in E. coli by competition or displacement (Brunel et al., 1995; Larios et al., 2002).
Another study which demonstrated repression of translation is by mechanism of entrapment seen during translation repression of α operon in E. coli. The α operon includes S4, S13, S11, the α subunit of RNA polymerase core enzyme and ribosomal protein L17. In this S4 acts as an allosteric effector and 11 regulates the S4 into two conformations, the active and the inactive form. The
30S subunit binds to both, however, only the active form can form a ternary
complex. The repressor and the 30S subunit bind the inactive form and are
trapped irreversibly into a dead end complex, which is stabilized by the protein
S4 inactive form (Schlax et al., 2001; Schlax et al., 2003).
In other regulatory mechanisms of translation by repression, studies have shown that ribosomal saturation can also lead to autogenous downregulation.
The ribosomes will compete and bind to their own mRNA IF3-L35-L20 operon, encoded by infC, rpmI and rplT respectively. IF3 negatively regulates infC by
binding its own mRNA. However, L20 binds to two regions, first one is a
pseudoknot formed by joining of the two nucleotide sequences that are at a
distance from each other (280 nt) and other site is formed into an irregular helix
by the two adjacent sequences, and both sites resemble (mimic) the original
binding site of L20 on 23S rRNA (Gullier et al., 2005; Gullier et al., 2005). L20
directly reduces translational efficiency of its own rpmI mRNA and also transmits
this negative effect to the downstream gene (rplt) in the operon through
translational coupling. Coupling mechanism was supported by the existence of
an inhibitory secondary structure (formed by 3’ terminal region of rpml, the rpml-
rplt spacer region and part of SD sequence of rplT) which prevents the
ribosomes to bind the RBS of rplT (Chiaruttini et al., 1996).
12 Regulation of translation under conditions of stress
In E. coli heat shock, cold shock, and other stresses like starvation, acid
pH, and high osmolarity induce translation of a global regulator rpoS encoded σS subunit of RNA polymerase (Brown and Elliot, 1996; Muffler et al., 1996). DnaK a heat shock protein also helps reducing degradation of σS levels by heat shock
and carbon starvation. The heat shock mediated protection to RpoS could be
provided by increased steady state levels of DnaK chaperone machines that
protect RpoS direct or indirectly from degradation (Muffler et al., 1997).
During the cold shock response the majority of the genes that regulate
transcription or translation stop being expressed, however a set of approximately
26 cold shock genes are expressed briefly. CspA is a cold shock protein that is
induced at lower temperatures and maintains efficient translation of mRNA at
lower temperatures by preventing the formation of secondary structures in RNA
(Jiang et al 1997). IraP a recently identified global regulator interferes with RssB
by direct protein-protein interaction, and prevents RpoS degradation (Bougdour,
et al., 2006).
However rpoS translation rates vary with different nutrient levels, under
ammonia starvation levels, rpoS translation increases slightly as compared to the
induction response under glucose and phosphate restriction. Glucose starvation
also induces translation of RpoS. Carbon starvation inhibits SpreE (RssB) a
response regulator via ClpXP protease mediated degradation of RpoS (Mandel 13 and Silhavy, 2005). In a similar context the genes that interplay during phosphate
starvation includes Pho regulon (PhoR) that is activated by low Pst levels, and
increases inorganic phosphate (Pi) levels by inducing gene expression and
providing the Pi by other pathways. PhoR activation induces the rpoS translation
during exponential phase along with Hfq as a chaperone protein and a small
RNA (Ruiz and Silhavy, 2003).
During nutritional deprivation states, E. coli adapts to the change by
slowing its overall metabolism and growth rate by immediately shifting into
stationary phase. Once the nutritional status is changed from poor to rich, many
physiological mechanisms are stimulated within the E. coli cell and the cell again goes back from stationary state into growing phase, Fis protein is involved in the adaptation to the nutrient shifts and BipA protein (a global regulator) regulates fis gene expression in a growth rate dependent manner by interacting with the 70S ribosome, induce conformational change which disrupts the interaction between
3’ end of 16S rRNA, and the 5’ UTR of fis mRNA.(Owens et al., 2004) Under decreased levels of carbon CsrA regulates translation of cstA gene by disrupting the binding of ribosomes (Dubey et al., 2003).
Small RNAs as regulators of translation
To date more that 200 small non-coding RNAs have been identified both in prokaryotes and eukaryotes, many more are being identified each year using computational and algorithmic approaches. Most of these small RNAs (>70)
14 identified so far have been characterized by Northern blotting. In E. coli, 30% of those identified, regulate gene expression at post-transcriptional level, usually by basepairing with their target mRNAs (Majdalani et al 2005).
Majority of these that have been characterized so far bind the RNA chaperone Hfq via AU-rich sequences (Storz 2004). Hfq aids in the basepairing of these small RNA with their specified targets by modulating the mRNA stability and translation (Majdalani et al 2005). Thus plays a crucial role for the regulatory mechanisms used by small ncRNAs OxyS, DsrA, RhyB, Spot 42, MicA, SgrS and many more on their mRNA targets rpoS, fhlA, sodB, sdhCDAB, galK, ompA, ptsG. These small RNA have been described in detail in the review of literature section on Hfq (Wasserman, 2002; Zhang et al., 2003; Majdalani et al., 2005;
Tjaden et al., 2006; Zhang et al., 2006; Ziolkowska et al 2006).
15 PART II
History of identification
Escherichia coli (E. coli) host factor (HF-I) protein (Hfq) is a nucleic acid binding protein that acts as a global regulator of gene expression by interacting
with small, non-coding RNAs. Hfq was identified in 1968 for the in vitro replication
of the (+) strand of phage Qβ RNA. Hfq binds with high affinity to single stranded
RNA, which implies that it may have a significant role in RNA metabolism
(Franze de Fernandez et al., 1968; Franze de Fernandez et al., 1972, Miranda et
al., 1997; Schuppli et al., 1997; Su et al., 1997). Homologs of Hfq have been in
more than 30 species of bacteria (Sun et al 2002).
Purification, sequencing, and crystal structure analysis
Purified Hfq protein from phage infected and uninfected E. coli cell occurs as a homo-hexameric, heat stable, small but abundant protein with102 amino acids long chain (Kajitani et al., 1994). Hfq protein is encoded by the hfq gene located at 94.8 min on the E. coli chromosome (Kajitani and Ishihama, 1991). A
single polypeptide chain has a molecular weight of ~12-13 kilo Dalton with an
apparent molecular weight of the active factor of ~ 70,000 to 80,000 (Franze de
Fernandez et al., 1972) that suggests the presence of subunit structure with 6
polypeptide chains.
16 Hfq is also defined as growth related protein. The intracellular level of Hfq ranges from 30,000 to 60,000 molecules per cell and increases with increase in the growth rate, and also in the exponential phase of cell growth. Largest fraction of the Hfq stays in association with the ribosomes in the cytoplasm whereas a small part is associated with the nucleoid (Kajitani et al., 1994; Azam et al.,
2000). In another study by (Tsui et al., 1994) an insertion mutation generated in the hfq gene resulted in decreased growth rate, increased cell length, osmosensitivity and also sensitivity to ultraviolet light, which is an indicator of a significant role of Hfq in E. coli cell physiology (Kajitani et al., 1994; Talukder et al., 2000).
Our present understanding of Hfq at the molecular level arises from its close resemblance to Sm and Sm-like protein in the N- terminal regions of approximately 80-120 base pairs. Recent data clearly indicated that Hfq a ring- shaped hexamer. This similarity was further supported by phylogenetic, biophysical, fold recognition, and homology modeling studies of E. coli Hfq
(Arluison et al., 2002; Sun et al., 2002; Moller et al., 2002; Zhang et al., 2002) and has been further confirmed by crystal structure analysis of Hfq hexamers from Staphylococcus aureus (Schumacher et al., 2002) E. coli (Sauter et al.,
2003) and Pseudomonas aeruginosa (Nikulin et al., 2005). On electron microscopic analysis of the crystal structure, the monomers of Hfq consist of five strongly bent, antiparallel β sheets capped by an N-terminal α helix and individual
17 subunits assemble into cyclic quaternary structures perforated by central cationic pores forming ring shaped homohexamers (Arluison et al., 2005).
Role as a chaperone protein
RNAs usually encounter two folding problems in the absence of proteins.
The first one arises primarily from alternate base pairings which then leads to the conformation of kinetically trapped inactive molecule. The second problem occurs due to the difficulty of some RNAs being nonspecific about their thermodynamically active tertiary structures which could be favored over other competing three dimensional structures (Herschlag, 1995). These folding problems could be solved with the help of two classes of RNA binding proteins and were defined as RNA chaperones. Group I RNA chaperones proteins are defined as proteins that bind nonspecifically to RNA sequence and prevent misfolding of RNAs or resolve the misfolded structures. In comparison, the other group of RNA chaperones includes those proteins that are able to specifically recognize the tertiary structures and stabilizing them (Herschlag, 1995).
Posttranscriptional and translational regulation
Hfq functions as a chaperone protein as well as a pleiotropic regulator that plays a role in the stability, replication, translation, and degradation of several target mRNAs. At its inception, Hfq was found to be required for Qβ replication of bacteriophage RNA (Franze de Fernandez et al., 1968 Su et al., 1997).
Previous work (Weber et al., 1972; Meyer et al., 1981) showed that replicase
18 holoenzyme binds to Qβ plus strand at two internal sites, S and M however these
interactions were detected only after initiation of synthesis. For the minus strand
synthesis to occur at 3’-end, Hfq was needed (Meyer et al., 1981; Barrera et al.,
1993; Miranda et al., 1997). Further studies proposed that binding of Hfq to the 3’ terminus of the phage sequence altered its tertiary structure and allowed for replication to occur (Miranda et al., 1997, Schuppli et al., 1997). Hfq also
increases access of the Qβ replicase to 3’ end of the positive strand, where
initiation of the negative strands synthesis occurs (Schuppli et al., 2000).Studies show that Hfq not only regulates the expression of genes at individual levels
(ompA or sodB), but control the gene expression of global regulator like Fur or σS
subunit of RNA polymerase. A study (Muffler et al., 1996) showed that Hfq was
required for the normal expression of the σS subunit of RNA polymerase and
regulates translation of rpoS. This subunit is an important regulator for
expression of many genes which are induced during entry into stationary phase
or in response to higher osmolarity. Hfq was also found to enhance rpoS translation by destabilizing inhibitory secondary structure present in the rpoS mRNA (Brown and Elliot., 1996).
Hfq negatively regulates the expression of mutS and mutH gene expression. MutS, MutL and MutH play a significant role in DNA repair pathways and stability of the chromosomes. Hfq affects the expression of mutS and miaA- hfq and its own in exponentially growing cells by making the transcripts unstable.
19 This involves an RpoS-independent mechanism and is consistent with a role of
Hfq as an RNA chaperone (Tsui et al., 1997).
Another example where Hfq acts as a chaperone is by destabilizing the
ompA mRNA. OmpA protein functions in the assembly and maintenance of the
outer membrane. Hfq regulates the stability of ompA message both in a growth
rate as well as growth phase dependent fashion. Previous studies using toe-
printing and in vitro translational assays on ompA showed that Hfq binds to the 5’ untranslated region (UTR) and interferes with the binding of 30S ribosomal unit to
5’ UTR to protect the ompA from cleavage by RNase E and the rate of degradation of ompA parallels the rate of growth and was detected to be more in slow growing cells, where the levels of Hfq were found to be more and negatively regulating the ompA mRNA levels (Vytvytska et al., 1998; Vytvytska et al., 2000).
OmpA mRNA was further used as a model to study and classify Hfq as an RNA chaperone by being dispensable after altering the structural stability of the message (Moll et al; 2003). Most recent studies demonstrated that Hfq controlled regulation of ompA mRNA is mediated by a small non coding RNA MicA
(previously SraD). In a toe printing assay it was established that upon binding to the translation initiator MicA blocks the access of the ribosomes in the stationary phase, facilitates RNase E cleavage and mRNA decay, and requires Hfq for the process to occur. The studies above establish two distinctive regulatory pathways of Hfq on mRNA, one being rate dependent and the other phase dependent (Rasmussen et al., 2005; Udekwu et al.,2005).
20 Other mRNA targets whose expression is under the direct control of Hfq
are those involved with the iron metabolism and include fur and sodB and
encoding the iron superoxide dismutase (Sakamoto and Touati, 1984). Hfq
inhibited translation of both fur and sodB mRNA as its direct effect (Dubrac and
2000; Dubrac and Touati, 2002). Studies performed by Masse and Gottesman
(2002) states that Fur represses transcription of RyhB RNA. RyhB repression in
turn favors the expression of sodB mRNA. Hfq regulates FeSOD synthesis via
Fur in a RyhB dependent mechanism. This study demonstrates both direct
(where Hfq exerts the effect itself) and indirect (involving Fur/RyhB) translational
control of Hfq (Vecerek et al., 2003). More recently in another study, it was
demonstrated by both in vivo and in vitro analysis that Hfq binds to AU rich
sequence on the sodB MRNA, this binding opens the internal loop and allows the
RyhB to bind which disrupts translation and stimulates the decay of both RyhB and sodB MRNA (Geissmann and Touati, 2004).
Another function of Hfq that characterizes it as a chaperone protein is its
role in enhancing and mediating RNA-RNA interaction, examples include Spot 42
RNA and galk mRNA, where a nucleoprotein complex is formed after Hfq
interacts with the Spot42 and its target mRNA facilitating the intermolecular
interaction by aligning the complementary sequences to its surface These
findings were observed on gel shift assays, co-immunoprecipitation assays, and
hydroxyl radical footprinting assays (Moller et al., 2002).
21 Interaction between OxyS RNA and fhlA mRNA is also mediated via Hfq
(Zhang et al., 1998; Zhang et al., 2002). OxyS RNA is an RNA that is induced under conditions of stress in E. coli. OxyS is defined as a global regulator and causes repression of some genes while activation of others. OxyS acts as a negative regulator of both fhlA mRNA and rpoS mRNA. In case of fhlA mRNA,
OxyS recognizes the two complementary basepair sequences in vitro, (Altuvia et
al., 1998; Argman and Altuvia, 2000) which were found to be stable even in the
absence of Hfq, however in vivo these conditions seemed to be less favorable, thus paving the way for Hfq to come in and enhance as well as stabilize the interaction. Coimmunoprecipitation and gel shift assays detected that Hfq was required to enhance the interaction. By RNase T2 footprinting, it was detected that Hfq changes the overall conformation of the stem and loops of OxyS RNA by cleaving them at some regions and protecting them at other sites making it more open (Zhang et al., 2002). OxyS also regulates rpoS and is induced by hydrogen peroxide.
RNA chaperone activity in vivo could be detected by measuring splicing efficiency in the T4 phage td intron by the protein StpA which had already shown
RNA chaperone activity in the in vitro assays (Clodi et al., 1999). In this study the researchers used the folding trap formed upstream of exon of the td gene and another at the 3’ of the intron and detected that in the absence of translation inserting one stop codon in the upstream exon was unfavorable for splicing, due to misfolding of pre-mRNA by exon-intron interaction (Semrad and Schroeder,
22 1998). StpA along with other RNA chaperone tested independently was able to
rescue the misfolded structures, giving them the right conformation that would
restore the splicing property (Clodi et al., 1999). Similarly using the group1 introns, Hfq was tested by both in vivo and modified in vitro toe printing assays to demonstrate rescuing of defective RNA folding trap in td gene of T4 bacteriophage into splicing efficient derivative and Hfq was able to do so, which again supports its role as an RNA chaperone (Moll et al., 2003).
Hfq has also shown an involvement in the polyadenylation of the mRNA.
To study this function of Hfq, the rpsO mRNA transcript was used as an example.
Various experimental data, both from in vivo and in vitro assays suggest that Hfq competes with polynucleotide phosphorylase (PNPase) polymerase and exonucleases (RNaseII and RNaseE) that have an affinity to bind to 3’ of the transcript and promote the decay. Hfq primarily blocks the access of exonucleases by binding to the 3’ end and simultaneously induces the lengthening of poly (A) tails catalyzed by poly(A) polymeraseI (PAPI). However, the 5’ end of the mRNA transcript was not involved. Further studies on Hfq mutants showed that only few were affected by the deletion, and could not bind and stimulate the length of poly(A) tails (Hajnsdorf and Regnier, 1999; Hajnsdorf and Regnier, 2000; Folichon et al., 2003; Mohanty et al., 2004; Folichon et al.,
2005; Ziolkowska et al 2006).
23 Hfq also indirectly controls the regulatory action of DsrA RNA on their
complementary target sequence by base pairing and is required for this function.
Until now out of the 220 bacterial genome sequenced, DsrA RNA has been identified in E. coli, Salmonella enterica and klebsiella pneumoniae (Rolle et al.,
2006). DsrA RNA is a small 85 nucleotide long, noncoding RNA and regulates
the expression of two global regulatory proteins, RpoS and HNS. DsrA binds by
base-pairing mechanism to the 5’ prime leader of rpoS MRNA. Like all other
RNAs DsrA RNA also requires interaction of Hfq to facilitate its action on rpoS
mRNA. Hfq enables the DsrA RNA to alter the conformation of rpoS mRNA
5’UTR, which further helps in the base-pairing between the two.
Assays demonstrate that the ternary complex formed between the three is
very brief, the site where Hfq binds on DsrA RNA is obscured after base-pairing
and Hfq is freed for recycling (Sledjeski et al 1996; Sledjeski et al., 2001; Brescia et al 2003; Lease and Woodson 2004). Additional findings from a recent study
(Koleva et al., 2006) suggest that S1 a large ribosomal protein also binds Hfq to interact with DsrA as well as rpoS mRNA and aids in DsrA mediated regulation of rpoS gene.
Mutational studies to characterize the binding role of Hfq
A study by Sonnleitner and his colleagues (2004) investigated the effect of
Hfq mutation on phage Qβ replication and demonstrated that the substitutions
24 that disrupted dodecamerization also inhibited Qβ replication. The other
parameter investigated was the effect of mutant Hfq on ompA translation, and
noted that Hfq mutant enhanced the translation when the deletion involved the residue between Hfq51-Hfq66, indicating that these amino acids (aa) are important
for binding of Hfq to small RNA. The group also studied the effects on DsrA
stability and demonstrated that Hfq51 does not bind DsrA as the stability was
reduced, however Hfq66 showed increased affinity to DsrA and enhanced its
stability presumably by inhibiting the action of RNase E. Role of PAP1 was also
studied and it established dsrA degradation did not occur in absence of PAP1,
which could be due to failure in the recruitment of PAP1 by Hfq66 mutant, thus
supporting the increased stability of dsrA in Hfq66.
Another study used site-directed mutagenesis D40, F42, K56 which
perturbed the RNA binding site surrounding the central pore of the
homohexameric ring, and had a mild effect on binding of A27 and rpoS MRNA,
while the double mutants D40 and F42 demonstrated wt like activity, however,
the DsrA binding was affected by K56A, this affect enabled us (Mickulecky et al.,
2004) to hypothesize more than one binding site on Hfq. Further mutations like
V43C did not affect the RNA binding, yet V43R did affect cell growth and rpoS
expression in presence of wt Hfq, reflecting the formation of heterohexamers,
with dominant negative function. A gene oppA encoding nutrient transporter and
periplasmic protein OppA is negatively regulated by ncRNA GvcB in presence of
Hfq; however V43R does not alter this ability. These two different effects of the
25 same mutant V43R, implicate several ways of binding of GcvB and DsrA RNA to different binding sites on Hfq protein (Zoilkowwska et al., 2006).
26 PART III
Discovery of RM systems
Restriction modification (RM) systems were identified over 50 years ago,
as the basis for either reducing or precluding phage infection. The need for this
defense against bacteriophage infection is widespread in the bacterial world, helping to shape the evolution (Price and Bickle, 1986).
Studies in the1950s revealed that phages infecting one strain poorly
infected another strain of the same species. The few productive infections that
did result yielded phage that had switched strain preference. This phenomenon
was termed host-controlled modification (Murray 2000).
The earliest examples of host-controlled modification turned out to involve
classical RM systems in E. coli K, and E. coli B (Bertani and Weigle, 1953). In
another study (Luria and Human, 1952) a nonclassical type of RM system was
involved. This ability to restrain invasion by phages helps to maintain the genetic
stability of bacterial populations by limiting immigration of foreign genetic material
(Kruger and Bickle, 1983).
Further analysis in the 1960s proved that the modification
methyltransferase, that methylates specific DNA bases, and the restriction
endonuclease, that cleaves the ummethylated DNA, recognize the same specific
27 DNA (Arber, 1965; Dussiox and Arber, 1965). Most restriction enzymes identified
so far have an accompanying methyltransferase component, and together form a
classical RM system (Murray, 2000). Experiments by Bertani and Weigle (1953)
with phage λ grown on E. coil C strain, E. coil B or E. coli K confirmed the
properties of the classical RM system.
Occurrence and functions of RM systems
Over 3800 RM systems have been identified so far, predominantly in
bacteria, though some have been seen in archaea and in viruses of certain
unicellular algae (Jeltsch, 2003). This distribution suggests that RM systems may have more than one function (Tock and Dryden, 2005). The prevailing view is that RM systems restrict phage infection. But RM systems limit this defense mechanism only to double stranded (ds) DNA templates, however RNA and single stranded (ss) DNA phages infect bacteria without being restricted efficiently or at all (Levin, 1993). In addition bacteriophages can overcome restriction by inhibiting the endonuclease activity, or by nucleotide base modification, and even phage coded methylation (Kruger and Bickle, 1983;
Kruger and Bickle, 1993; Tock and Dryden, 2005). Other possible roles for RM systems include DNA recombination and repair (Tock and Dryden, 2005). RM systems may also behave as selfish genetic elements by post-segregational killing (Kusano et al., 1995; Nakayama, 1998; Kobayashi, 2001).
28 Classification of RM systems
RM systems are classified according to the enzyme composition, cofactor
requirements, recognition sequence specificity, cleavage sites, and number and
organization of subunits. Four types of RM systems and several subtypes have
been characterized to date. My research has explored the regulation of one type;
this background illustrates the remaining territory to be explored.
Type I RM system
Type I RM systems are present extensively in prokaryotes and about 600
enzymes have been identified to date. Out of these 69 restriction genes, 35
methyltransferase genes, and 57 specificity subunits have been characterized
biochemically representing under a third of the total number (Roberts et al.,
2007). Type I R-M systems are the most complex system, found in many
bacteria. They are hetero-oligomers, composed of two REase subunits (R) that
cleave the DNA, a single specificity subunit (S) that determines the DNA
sequence recognized, and two MTase subunits (M) required for methylation with
S-Adenosylmethionine (Adomet) as the methyl donor. The methyltransferase
activity methylates the N6 position of adenine, with a predilection for
hemimethylated sites that is not frequent with prokaryotic MTases. REase activity
requires ATP, Adomet and Mg2+ (Kruger and Bickle, 1993; Murray, 2000;
Bujnicki, 2001; Roberts, et al., 2003; Tock and Dryden, 2005).
29 When the REase component contacts an unmethylated target sequence,
ATP-dependent translocation of DNA is initiated from both directions, resulting in
supercoiling of DNA. Cleavage occurs at a site that is far from the target recognition site and is driven by stalled translocation. Collision between the two translocating complexes leads to cleavage at both strands of DNA. EcoKI and
EcoBI are the best known examples of type I RM system and both complement each other, with the ability of EcoKI to limit phage propagation by factors of between 103 and 108 (Webb, et al., 1996). The type I RM system has been further categorized into four (A, B, C and D) subtypes based on their complementation properties (Kruger and Bickel, 1993; Murray, 2000; Bujnicki,
2001; Roberts, et al., 2003; Tock and Dryden, 2005).
Type III RM system
This system is similar to Type I RM systems, but less intricate. Enzymes in
this group are hetero-oligomers, composed of two polypeptides, the REase (res)
and MTase (mod) subunits. The MTase methylates the N6 position of adenine
but modifies only one strand. It functions independently and utilizes Adomet as
methyl donor forming hemi-methylated sequences. This partial modification is not
lethal because cleavage always ensues on an unmethylated sequence thus
hemi-methylation protects the DNA (Wilson, 1991; Roberts, et al., 2003; Tock
and Dryden, 2005).
30 However, the REase component cannot function independently. It needs the MTase in order to initiate cleavage and also requires Mg2+ and ATP. For the
cleavage to occur, this enzyme system needs two copies of an asymmetric recognition sequence in an inverse orientation and within the DNA molecule.
During translocation the res-mod complex stays in contact with the recognition sequence and cleavage begins either by collision of res-mod complex or by the stalled translocation at a site that is 25-27 bases away from one of the two copies of the recognition site. Type III RM systems are present in phages, and Gram negative bacteria. Eco1PI and Ecop151 are two well characterized examples and so far about 160 putative type III RM systems have been identified by sequencing prokaryotic genome (Wilson, 1991; Roberts, et al., 2003; Tock and
Dryden, 2005).
Type IV RM system
This system includes enzymes with one or two REase components that only cleave sequences that have been methylated, hydroxymethylated or glycosyl- hydroxymethylated. Mg2+ and GTP are used for translocation and cleavage which
is initiated by stalled translocation (Pieper, et al 1997). McrBC from E. coli K12 has been studied for the cleavage pattern, and the enzyme identifies two copies of the dinucleotide sequence consisting of a purine followed by a methylated cytosine (at N4 or C5 position) which are separated by a length ranging from 40-
3000 bp. Cleavage occurs at a site 30 bp away from one of the recognition strands (Sutherland, et al., 1992).
31 Type II RM systems
These are the most abundant and more than 3600 type II RM systems
have been identified so far representing nearly half of the total number. This is
also the type upon which part of thesis focuses. It has a simple subunit
organization which differentiates it from the types I, III and IV. In most type II RM
systems, REase and MTase function independently. Mg2+ alone is required for
REase activity and Adomet alone for the MTase activity. Both the REase and
MTase recognize the same target sequence, which is usually a short palindrome
of 4-8 bps length. Type II REases are generally homodimers that cleave at or
near the specific DNA sequence, producing two strands with 5’ PO4 and 3’ OH
end. Type II MTases usually act as monomers to modify a specific base, cytosine
at N4 or C5 position or adenine at N6, on each strand of the duplex. Owing to their
sequence-specific function, type II RM systems have become indispensable tools of modern biological technology. Some type II RM systems recognize interrupted palindromes that include unspecified sequences (Kruger and Bickle, 1993;
Murray, 2000; Bujnicki, 2001; Roberts, et al., 2003; Tock and Dryden, 2005;
Pingoud et al., 2005).
Subtypes of Type IIRM system: IIP, IIA, IIB, IIC, IIE, IIF, IIG, IIH, IIM, IIS, and
IIT
Type IIP is a representation for the orthodox type II RM system, where the
REases are homodimers that recognize symmetrical palindromes, and cleave at
fixed symmetrical location which lies either within or adjoining the recognized
32 sequence. The best analyzed examples include EcoRI and EcoRV (Hedgpeth, et al., 1972; Schildkraut I et al., 1984 Roberts, et al., 2003). This also includes the
PvuII system that I have studied.
Type IIA REases recognize asymmetric sequences. Enzymes in this group are useful for production of nicking enzymes. Normally enzymes from this group consist of one REase gene and two MTase genes for modification of each strand of the asymmetrical sequence. Example includes Bpu101, which is hetero-dimer with two R genes, and FokI with fused R and M genes (Roberts, et al., 2003; Stankevicius, et al., 1998).
Type IIB cleaves DNA on both sides of the target sequence, and on both strands. For example BplI cleaves the top strand 8 nucleotides before and 13 nucleotides after the recognition sequence, while the bottom strand is cleaved 13 nucleotides before and 8 nucleotides after the gapped recognition sequence
(Roberts, et al., 2003; Vitkute et al., 1997).
Type IIC overlaps other subtypes. It includes all cases where both REase and MTase activities occur on a single polypeptide chain. This includes all the enzymes under subtype IIB; IIG and some from Type IIH. BcgI as one of the earliest identified example. It has both REase and MTase on one subunit and target recognition domain on a second subunit, while others like Cjel and HaeI
33 have all three functional domains on single chain (Roberts, et al., 2003; Kong, et al., 1994; Kong, 1998).
Type IIE enzymes use two copies of their recognition sequence for effective cleavage, one site being used as the recognition site for cleavage and the other acting as an allosteric effector. EcoRII and NaeI are the two enzymes studied in detail. Removal of the effector domain from EcoRII converts it into
Type IIP. Sau3AI dimerizes in presence of DNA and behaves like Type IIE having both allosteric as well as catalytic site (Roberts, et al., 2003; Friedhoff et al., 2001).
Type IIF enzymes present most often as homotetrameric structures, and need two copies of target sequence for efficient cleavage. Examples include
CfrIOI, NgoNIV, SfiI and SgrAI, which is a dimmer in solution but functions as a tetramer upon binding to DNA (Roberts, et al., 2003; Daniels, et al., 2003).
Type IIG enzymes behave like Type IIC from all aspects, having both
REase and MTase domains exist on a single polypeptide chain, and requiring
Adomet for stimulation. They may recognize both symmetric and asymmetric sequences, such that Type IIP and Type IIA may belong to Type IIG. Most Type
IIG including Eco57I show characteristics that could serve as useful engineering tools (Roberts, et al., 2003; Rimseliene, et al., 2003).
34 Type IIH is genetically identical to Type I RM system and functionally it behaves like Type II RM systems. In this group AhdI enzyme has been characterized so far and recognizes a gapped target sequence (Roberts, et al.,
2003; Marks et al., 2003).
Type IIM enzymes recognize a specific methylated sequence and cleave the DNA at fixed site. DpnI has been characterized so far (Roberts, et al., 2003;
Herman and Jeltsch, 2003).
Type IIS enzymes behave like Type IIA, where one of the strands involved in cleavage lies outside the limits of recognized target DNA sequence. Two domains exist in this subtype, one for target site recognition and the other for catalysis, which also serves as a homodimerization domain. FokI is an example of Type IIS that uses two recognition sites before cleaving DNA. Type IIS enzymes have been used for generating rare restriction sites (‘Achilles’ heel cleavage), chimeric nucleases and strand-specific endonucleases (Roberts, et al., 2003; Koob et al., 1988; Koob et al., 1988; Chandrasegaran and Smith 1999;
Kandavelou, et al., 2004).
Type IIT enzymes are heterodimeric, with many of these being used to create nicking enzymes (cleaving one strand only) like BbvCI, BpuI0I and BslI (Roberts, et al., 2003; Zhu, et al., 2004).
35 Regulatory mechanisms in the RM system
In Type II RM systems, the genes for REase and MTase present either in
the same orientation or in the opposing orientation that is either convergent or
divergent, having either R or M upstream along the sequence. In the Type II RM
systems where the genes showed opposite orientation, it was proposed that their
regulation occurred from two independent promoters as was also true for the
Type I RM system. However, in those Type II RM systems where both genes displayed the same orientation, it was proposed that owing to the close proximity of the genes, regulatory control operated from a single promoter.
The control of relative activities of MTase and REase is critical because a reduced ratio of MTase to REase would lead to cell death via autorestriction (De
Backer and Colson, 1991), while too high of a ratio would fail to protect against the invading phage DNA. Furthermore many RM systems move from one host to another either by residing on a plasmid or by transduction, transformation, or Hfr- type conjugation. The new host’s DNA requires fairly complete modification
before the restriction endonuclease activity appears, thus the need for tight
regulation of genes for RM systems is obvious (Vijesurier, et al., 2000).
To date, three mechanisms have been described. One of them employs the HTH motif identified at the N-terminus of the methyltransferase. Binding of
HTH motif to the promoter can lead to repression of initial high expression of
MTase gene. In SsoII and EcoRII RM systems MTases acts as transcriptional
36 regulators and maintain a coordinated expression of restriction and modification
enzymes (Som and Friedman, 1993; Butler and Fitzgerald, 2001). However this
would not explain delayed REase expression.
A second type of regulatory mechanism of RM system was based on the
coordinated activity of the two promoters (R and M) with an overlap region. The
overlap region includes a unique DNA target sequence whose methylation status
directs the efficiency of expression of the two promoters (Beletskaya, et al.,
2000).
In SalI RM system the gene operon is mainly transcribed from a promoter
located upstream from salIR, the first gene of the operon and at the 3′ end of
salIR, there is a second promoter that allows independent transcription of the
MTase gene (Alvarez, et al 1995). In EcoRI also the R and M gene constitute an
operon. The promoter which regulates their expression is present upstream of
the R gene, in addition there is a second promoter which is specifically meant for
M gene expression is positioned within the R gene (Rosenberg et al., 1981;
O’Connor and Humphreys 1982). It is proposed that M gene specific promoter
should allow the M gene expression in the absence of R gene that might play a
role in sequential expression of MTase activity and REase activity upon the entry
of EcoRI gene system into a new host cell (Greene et al., 1981).
37 Regulation by C proteins
A third mechanism for controlling RM involves a specialized small protein,
designated as controller (C) protein. In a subset of type II RM systems an open
reading frame (ORF) approximately 100 codons preceded the R gene (Tao et al.,
1991) and the R gene and M gene demonstrated different orientation. This subset includes RM systems such as PvuII (Vijesurier, et al., 2000) BamHI (Ives, et al., 1992), BglII (Anton et al., 1997), Eco721 (Rimseliene, et al., 1995), EcoRV
(Zheleznaya, et al., 2003), Esp13961 (Cesnaviciene, et al., 2003) and SmaI
(Heidmann, et al., 1989) which all depend on C proteins (Tao et al., 1991) for coordinated expression.
The small ORF that precedes the R gene (controller) protein and in some cases partially overlaps the R gene is named the C−protein and it plays an
important regulatory role (Tao and Blumenthal 1992). Genes coding for C protein
bind to C boxes and activate the expression of their own gene and the
downstream endonuclease (Bart, et al., 1999).
The small ORF in pvuIIC and bamHI is between the two other genes of the PvuII RM and BamHI RM system. Both C.PvuII and C.BamHI protein are required for expression of themselves and of the downstream REase, which delays the appearance of nuclease activity following the transfer of the RM system into the new host. By the time the C protein accumulates the MTase has had time to modify DNA sufficiently to overcome the deleterious action of REase
38 (Brooks et al., 1991; Ives et al., 1992; Nathan and Brooks, 1988; Tao and
Blumenthal 1992; Tao et al., 1991).
In both the PvuII RM and BamHI RM systems the C protein stimulated the expression of REase. In the PvuII system, the opposing pvuIIC and pvuIIM transcripts overlap by 60 nt at the 5’ end; their hybridization is suggestive of other regulatory roles (Tao et al 1991, Tao and Blumenthal 1992; Anton et al., 1997;
Brooks et al., 1991; Ives et al., 1992; 1995; Nakayama and Kobayashi, 1998;
Rimseliene et al., 1995; Vijesurier et al., 2000).
Analysis of the pvuIIC sequence predicted a helix-turn-helix (HTH) structure, similar to many other known activators or repressors of gene expression (Wintgens and Rooman, 1996). Our laboratory has identified more than 20 C ORFs from a broad range of bacteria, all showing a ~10 kDa subunit
HTH protein lying upstream of and in the same orientation as that of the endonuclease gene (Tao and Blumenthal, 1992; Knowle et al., 2005). Since
C.PvuII, C.SmaI and C.BamHI display substantial sequence similarity, they were tested for cross-complementation, and it was demonstrated that C-proteins from
PvuII, and SmaI could reestablish restriction in a bamHIC mutant but not as efficiently as C. BamHI did in a pvuIIC mutant (Ives, et al., 1995).
39 Crystal structure of C proteins
X-ray diffraction analysis of C.AhdI, an ortholog of C.PvuII revealed an all-
α protein that includes a classical HTH domain and belongs to the Xre family of
transcriptional regulators. Dynamic light scattering and crystal structure analysis
indicate that C.AhdI forms a homodimer, with low dimer stability that may
significantly contribute to the “all or none” behavior of the expression that delays
the transcription of the C and R genes (McGeehan, et al., 2004; McGeehan, et
al., 2005).
Another group discovered C.BcII while they were investigating expression
of the BcII RM system. Their work revealed negative regulation of the M.BcII in
the presence of C.BcII. They also characterized the crystal structure of the
C.BcII, consisting of five α helices: two of these form the HTH motif and the
remaining three form a dimer interface. Structure of C.AhdI is highly similar to
that of C.BcII (Sawaya, et al., 2005).
Further study on C.AhdI led to a proposed a mechanism for both positive
and negative regulation of endonuclease expression. It is proposed that C.AhdI
at moderate levels interacts with RNAP and stimulates transcription. As the
levels increase further, a second dimer competes with RNAP for binding,
reducing transcription of its own gene as well as that of the endonuclease gene
(McGeehan, et al., 2006).
40 MANUSRIPT I: Published paper
Escherichia coli Hfq has distinct interaction surfaces for DsrA, rpoS and poly(A) RNAs
Peter J Mikulecky1, Meenakshi K Kaw2, Cristin C Brescia2, Jennifer C Takach1, Darren D Sledjeski2 & Andrew L Feig1
1 Department of Chemistry, Indiana University, 800 E. Kirkwood Avenue, Bloomington, Indiana 47405, USA.
2 Department of Microbiology and Immunology, Medical College of Ohio, Toledo, 3055 Arlington Avenue, Ohio 43614-5806, USA.
Nature Structural & Molecular Biology 11, 1206 - 1214 (2004) Published online: 7 November 2004; | doi: 10.1038/nsmb858
41 ABSTRACT
The bacterial Sm-like protein Hfq facilitates RNA-RNA interactions involved in post-transcriptional regulation of the stress response. Specifically, Hfq helps pair noncoding RNAs (ncRNAs) with complementary regions of target
mRNAs. To probe the mechanism of this pairing, we generated a series of Hfq
mutants and measured their affinity for RNAs like those with which Hfq must
associate in vivo. We tested the mutants' DsrA-dependent activation of rpoS, and their ability to stabilize DsrA ncRNA against degradation in vivo. Our results suggest that Hfq has two independent RNA-binding surfaces. In addition to a well-known site around the core of the Hfq hexamer, we observe interactions with the distal face of Hfq, a new locus with which mRNAs and poly (A) sequences associate. Our model explains how Hfq can simultaneously bind a ncRNA and its mRNA target to facilitate the strand displacement reaction required for Hfq- dependent translational regulation.
42 INTRODUCTION
Hfq protein from Escherichia coli was first described in connection with Q
-phage replication1, 2. Hfq has recently emerged as a central player in post-
transcriptional gene regulation as mediated by bacterial ncRNAs3, 4, 5, 6.
Escherichia coli Hfq mutants show disrupted signaling in stress response
pathways7, 8, arising from the need for Hfq to mediate base-pairing between
regulatory ncRNAs and their mRNA targets. Examples of these partnerships
include DsrA-rpoS 7, 9, 10, OxyS-fhlA 11, 12, OxyS-rpoS 13, RprA-rpoS 14, RyhB- sodB 15, 16, 17 and Spot42-galETKM18.
Complexes between ncRNAs and their mRNA targets function in several
ways. Most commonly, complexed structures lead to translational activation or
repression by remodeling mRNA regulatory regions containing the ribosome-
binding site (RBS) and/or start codon. Alternatively, the interaction can enhance
decay of the target mRNA16 or simply block translation11. Clearly, Hfq facilitates
base-pairing between ncRNAs and their targets, but how it does so is poorly
understood. How the chaperone function relates to other Hfq activities such as
the control of poly(A) tail elongation19, 20 and regulation of mRNA stability21, 22 is
also unknown.
Hfq shares sequence similarity to the eukaryotic Lsm proteins23, 24, 25, 26, 27.
In the Conserved Domain Database28 Hfq is listed under the Sm and Sm-like
43 protein family as well as among the eubacterial Hfqs. An alignment of the
conserved Lsm and Hfq motifs shows that the Sm1 and Sm2 regions overlap
with the Hfq motif (Fig. 1). Crystallographic characterization of Hfq revealed a
classical Sm fold, as predicted from sequence alignment and homology
modeling27, 29. Whereas eukaryotic Sm proteins form heteroheptameric rings30, 31,
32, Hfq forms homohexamers similar to the archaeal Sm proteins33, 34. RNA-
binding contacts are observed for both Hfq and Sm proteins in cocrystal
structures with short (A+U)-rich oligonucleotides, and show that these small RNA
substrates interact with their protein partners in a similar manner (Fig. 1).
Two parallel but nonexclusive models have been proposed to explain how
Hfq promotes intermolecular base-pairing. In the first model, Hfq acts explicitly as
an RNA chaperone, partially unfolding one or both RNAs35, 36. In the second
model, Hfq binds both RNAs, increasing their local concentration to induce the
interaction. Several studies have probed RNA structural changes upon Hfq
binding. DsrA ncRNA showed no substantial secondary structure changes37. The rpoS mRNA 5' untranslated region (5' UTR) was not assayed for structural changes in this study, but more recent work indicates that the rpoS leader sequence also remains unchanged upon binding Hfq (R. Lease and S. Woodson,
Johns Hopkins University, personal communication).
Several studies suggested that more than one RNA can simultaneously assemble onto Hfq. Work by two groups has recently shown that stable ternary
44 complexes containing DsrA, rpoS mRNA and Hfq can form (P.J.M., E. Espinosa,
Indiana University, and A.L.F., unpublished data; R. Lease and S. Woodson,
Johns Hopkins University, personal communication). These complexes represent product-like states wherein the two RNAs are already base-paired with one
another on Hfq. Similar ternary complexes have been observed in
coimmunoprecipitation experiments25. It seems possible, therefore, that the
single RNA-binding site observed crystallographically is not sufficient for Hfq's
ability to promote intermolecular base-pairing.
We addressed these questions through a mutational analysis of Hfq,
probing in vitro binding to several model RNAs that represent species with which
Hfq must interact. Hfq mutants were assayed in vivo using a reporter assay and
RNA lifetime experiments. Together, the results support a model wherein at least
two independent RNA-binding sites exist on the Hfq hexamer, and juxtaposition
of bound RNAs facilitates base-pairing.
45 RESULTS
Hfq mutagenesis
To identify amino acids essential for RNA binding, we constructed a series of E.
coli Hfq missense mutants (Fig. 1). Hfq Y55A, Hfq K56A and Hfq H57A contain
mutations that cluster around the central cavity of the hexamer, a region that
interacts with short (A+U)-rich sequences in the Hfq−RNA cocrystal structure29.
Other mutants, such as Hfq D40A, Hfq Q41A and Hfq F42A lie along the
proximal surface of the torus. Minimal binding site studies have shown that Hfq
preferentially binds (A+U)-rich sequences adjacent to double-stranded regions37,
38. Thus, these sites represent a potential contact surface for the duplex were it to lay down onto Hfq adjacent to the site at which the (A+U)-rich sequence binds.
Hfq Q8A and Hfq D9A represent a series of highly conserved residues that lie on the proximal face of the torus that might contact RNAs a bit farther from the
central cavity. Finally, the Hfq Y25A, Hfq Y25N, Hfq Y25D, Hfq I30D, Hfq Q53A
and Hfq S60A mutations represent sites on the distal face of the Hfq hexamer. All
mutants were expressed as His6-tag fusion proteins. In control experiments, the
His6-tag had no substantial effect on the binding properties of Hfq in vitro (as
assayed by gel shift and calorimetry) and His6-tagged Hfq fully complements an
hfq- strain in vivo.
RNA-binding properties of mutant Hfqs
Previous data from our labs suggested that two or more RNA-binding surfaces
46 might be present on Hfq37. When poly(A) was used in an attempt to compete away DsrA in a gel shift assay, a supershift was observed rather than competitive
binding37. Therefore, to assess the effect of these mutations on RNA binding,
several different RNA substrates were tested, including DsrA, RNA-U (a 7-
nucleotide (nt) RNA with the sequence AU5G), rpoS mRNA 5' UTR and A27 (a 27- nt synthetic poly(A) oligomer). DsrA and RNA-U represent the ncRNAs, all of
which contain (A+U)-rich sequences as part of their minimal binding element.
This comparison is reinforced by the fact that short poly(U) RNAs have been
shown to supplant DsrA in competitive binding experiments37. rpoS mRNA 5'
UTR and A27 represent the target mRNAs regulated by the ncRNA, corresponding to the 5' and 3' termini of the mRNA, respectively.
RNA binding was probed using gel mobility shift assays and isothermal titration
calorimetry. In the case of DsrA, the RNA shifts, and then supershifts, as it binds first one then a second equivalent of Hfq hexamer (Fig. 2a). The rpoS mRNA 5'
UTR construct that we used also probably binds at least two equivalents of the
Hfq hexamer. Because the initial species never accumulates significantly we have treated the rpoS mRNA-binding data as a single transition providing an apparent K d (Fig. 2b).
We consolidated the data from the 4 RNA substrates for each mutant Hfq,
represented as the G relative to binding wild-type Hfq (Fig. 2c). DsrA binds to
wild-type Hfq with an affinity of 21 1 nM (reported as hexamer, 126 nM
47 monomer). Under the same conditions, the second binding event occurs with a K
d of 94 nM (reported as hexamer, 564 nM monomer). This value is within two-fold
of those previously determined37. DsrA affinity is affected by residual RNA often
found to copurify with Hfq. The RNase A treatment in the current purification
significantly reduced residual RNA and is probably the origin of the tighter
affinities measured here.
Most of the mutations had relatively small effects on DsrA-binding affinity (Table
1), the largest being on the order of 20-fold decreased affinity, corresponding to a
G of 1.8 kcal mol-1. The modest effects of the point mutants could have
resulted from polyvalent interaction between the larger RNAs and Hfq masking
defects in one of the binding domains37. If this were true, the smaller RNAs
(RNA-U and A27) would provide a more sensitive gauge of whether the residue
participates in the RNA-protein interaction.
Under the same gel conditions, RNA-U exhibits poor affinity for the wild-type Hfq
hexamer (2.5 0.2 M hexamer, 15 M monomer). This result is consistent with minimal binding studies that have shown Hfq preferentially binds (A+U)-rich sequences adjacent to base-paired elements37. Despite the weak interactions,
binding is perturbed by specific mutations. Hfq D9A showed eight-fold tighter
binding to RNA-U. Mutant proteins Hfq Y55A, Hfq Y55W, Hfq K56A and Hfq
H57A were sufficiently defective that no binding was observed, even at the
highest protein concentrations.
48 Because RNA-U bound so weakly, we compared its affinity to those of a pair of
RNAs that should not specifically bind Hfq—the substrate and ribozyme strands
of hammerhead ribozyme 16 (ref. 39). These RNAs showed no gel shift behavior
up to 30 M Hfq monomer. The interaction of RNA-U with the mutant Hfqs clearly
approaches that of pure nonspecific binding under these conditions. At the same time, the deleterious effects of mutations within the Sm2 core motif suggest that
RNA-U binding occurs at the crystallographically observed site.
With respect to rpoS mRNA 5' UTR binding, the collection of mutations we prepared showed few effects. The largest defect observed across the series, with
Hfq K56A, was only two-fold. This result indicates that perhaps not all RNAs bind via the central core and the Sm2 motif. Distal face mutations also did not significantly affect rpoS mRNA 5'-UTR binding. Two double mutants (Hfq Y25D
Y55A and Hfq I30D Y55A) that combine defects in the Sm2 motif with those on
the distal face show reduced affinity for rpoS mRNA 5' UTR, however these data
suggest that rpoS mRNA 5' UTR may interact with both faces of Hfq.
Very different results were observed in the case of A27 binding. Gel shift
experiments measured an affinity of 39 1 nM for wild-type Hfq hexamer (234 nM in monomer). Of the proximal face mutants, the most impaired was Hfq
Y55A, but this protein had only a three-fold effect on binding. These results suggest that the Sm2 region at the center of the torus does not interact in any significant way with A27 and is consistent with the additive binding behavior
49 previously observed between DsrA domain II and poly(A)37. The distal face
mutants showed different behavior, however. Mutations at Tyr25 and Ile30
affected A27 binding, leading to a ten-fold loss in affinity for Hfq Y25D and Hfq
I30D. Double mutants showed no additional changes in affinity for A27 when Sm2
motif changes were combined with distal face mutations. We therefore infer that
poly(A) interacts with the distal face of Hfq.
Competition experiments reveal two independent binding sites
Previous experiments with wild-type Hfq showed what seemed to be mutual
37 binding of A27 and DsrA to Hfq . In light of the results described above, we
carried out a series of competition experiments using gel shift assays to look at the effect of A27 addition to binary complexes containing either DsrA or rpoS
mRNA 5' UTR prebound to Hfq (Fig. 3 and Table 2). We observed markedly
different behavior for wild-type Hfq, Hfq Y25D and Hfq K56A. Concomitant
binding of A27 and either DsrA or rpoS mRNA 5' UTR was observed at
concentrations of wild-type Hfq sufficient to promote the formation of the
37 DsrA−(Hfq6)2 species, consistent with our previous findings . When the distal
face mutant Hfq Y25D was used, additive binding was completely abolished, indicating that A27 binding was independent of DsrA and rpoS mRNA 5' UTR
(data not shown). When the Sm2 mutant Hfq K56A was used, a third outcome
resulted. The addition of A27 displaced DsrA; it similarly displaced rpoS mRNA 5'
UTR, albeit less efficiently. The band resulting from DsrA displacement by A27
50 migrated more slowly in the native gel than unbound DsrA, with mobility similar to
that of DsrA dimer (data not shown).
The competition experiments lead to two conclusions. First, Hfq K56A binds DsrA
improperly, allowing for competition by A27. Second, this abnormal binding mode possibly leads to altered folding of the ncRNA and assembly of DsrA dimers—a trait that could lead to adverse consequences in vivo.
Isothermal titration calorimetry
An alternative way to measure Hfq-RNA interactions is with isothermal titration calorimetry (ITC)40, 41, 42. This method offers several advantages over the gel shift experiments. Reaction stoichiometry is more readily determined from the data.
Furthermore, detailed thermodynamic information is obtained more accurately
than from electrophoretic methods, because the enthalpic contribution of the free
energy is measured directly.
A typical ITC experiment is shown in Supplementary Figure 1 online. Consistent
with the gel shift experiments, DsrA, rpoS mRNA 5' UTR and poly(A) showed
tight binding to Hfq hexamer with K d values in the low nanomolar range.
Because of the weak interaction between Hfq and RNA-U ( 2.5 M from gel shift
analysis), we could not achieve the necessary sample concentrations to study its
binding accurately using this method. Hfq Y55A showed a 50-fold reduction in
binding affinity whereas the Hfq K56A showed a 30-fold reduction in binding
affinity for DsrA, whereas Hfq H57A had nearly wild-type affinity. Both results are
51 consistent with the gel shift experiments (Supplementary Table 1 online).
The apparent stoichiometries from the ITC experiments are consistent with Hfq
acting as a preformed hexamer and binding a single equivalent of DsrA or rpoS
mRNA 5' UTR. Thus, the ITC data imply that the binding of a second equivalent
of Hfq to an RNA (K 2) may result from working under trace RNA conditions, as
typically done in gel shift assays. An unexpected result was observed in poly(A)
experiments, however. Fitting of the ITC data suggests that A18 (a shorter, 18-nt
poly(A) RNA was used in these experiments to diminish the likelihood that
multiple Hfqs might bind to the longer RNA43) binds two identical sites per Hfq
hexamer.
Binding enthalpies ( H°) of -40 to -80 kcal mol-1 were measured for the same three substrates with corresponding binding entropies ( S°) of -100 to -250 cal
mol-1 K-1 (Supplementary Table 1 online). These values show that the enthalpy of
the interaction is quite favorable, but largely offset by highly unfavorable entropy.
As these parameters combine the energetic contribution of the binding
phenomenon with any potential RNA rearrangements, a detailed interpretation
requires further study.
Accumulation and complementation analysis of Hfq mutants
Within a living cell, the interaction of Hfq with mRNAs and ncRNAs is much more
complex than that probed with our in vitro model system. To determine whether
52 the Hfq mutants function in vivo we used an rpoS::lacZ reporter system in an hfq- strain of E. coli (Fig. 4)7. Expression of this reporter requires both transcription
and translation of rpoS 7. The assay tests for two phenomena. First, plasmid-
borne mutant Hfqs must accumulate, and second, they must facilitate proper
post-transcriptional regulation of rpoS. Because DsrA is the predominant ncRNA
involved in the translational regulation of rpoS at low temperatures, all assays
were conducted at 30 °C (ref. 44). To prevent overexpression of Hfq, the mutants
were expressed under control of the inducible araBAD promoter45. In agreement
with previous work7, in the absence of arabinose or with vector alone, very little
Hfq accumulated (Fig. 4b) and RpoS::LacZ expression was low (Fig. 4c).
Addition of 150 M arabinose induced wild-type expression levels of Hfq and
normal translation of RpoS::LacZ (Fig. 4c).
As misfolded proteins tend to be degraded and cleared from the cell, western
blotting was used to ensure that mutant Hfqs accumulated normally at the same
arabinose concentration. Most of the mutants provided normal expression under
these conditions, as can be seen for representative mutants like Hfq Y55A and
Hfq K56A. Hfq D40A and Hfq F42A (Fig. 4b, lanes 6 and 7) showed abnormal
accumulation, however. Circular dichroism (CD) and dynamic light scattering
verified that these proteins fold and hexamerize in vitro, but they may be
sufficiently destabilized to pose a problem in vivo. Because there was a
significant decrease in the Hfq concentration in these cells, any loss of activity in
53 the reporter complementation assay (Fig. 4c) could result from either defective
RNA binding or decreased protein accumulation.
As expected from the in vitro experiments, Hfqs containing mutations in the Sm2 motif, including Hfq Y55A and Hfq K56A, did not complement the hfq- phenotype.
As these proteins were present at wild-type levels based on western blot analysis, these results suggest that loss of in vivo activity for these mutants was not due to decreased accumulation. Notably, Hfq H57A and Hfq Y55W, mutants that do not bind RNA-U but retain wild-type DsrA and rpoS mRNA 5' UTR affinity in vitro, retained their in vivo activity. This finding suggests that the small changes in DsrA binding observed in vitro for Hfq Y55A and Hfq K56A are more diagnostic of the in vivo behavior than RNA-U binding. Other mutations within the phylogenetically conserved Sm2 region showed no defects in our assay.
Assessment of DsrA stability in vivo
RNA lifetime analysis has been used as another measure of Hfq activity in vivo 7,
16, 38. In the presence of Hfq, DsrA has been shown to degrade significantly more
slowly than when Hfq is absent7. We measured DsrA lifetimes for both Hfq
mutants that had significantly reduced activity in complementation assays that
could not be explained by simple accumulation defects. Hfq Y55A and Hfq K56A
show markedly different behavior after inhibition of transcription with rifampicin
(Fig. 5). Whereas Hfq Y55A led to long-lived RNA similar to wild-type Hfq, in the
presence of Hfq K56A, DsrA degraded rapidly. The behavior occurred with
54 biphasic kinetics. DsrA degraded in the fast phase, accounting for 90% of the total, was less stable than in the absence of Hfq entirely. The remaining 10% of the RNA was reasonably long-lived.
55 DISCUSSION
Site-directed mutagenesis was used to probe the interaction between Hfq and
four RNAs that represent some of the ncRNAs and mRNAs with which Hfq
interacts in vivo. The effects of these mutations were assayed both in vitro and in vivo. Sites of mutation were chosen based on structural and phylogenetic information. Structurally, we can group the mutants into three categories: those
affecting the central cavity (Y55A, K56A and H57A), the proximal face (Q8A,
D9A, D40A, Q41A, F42A and I59A) or the distal face (Y25X, I30X, Q53A and
S60A). Notably, mutations at these loci had distinct effects on behavior, both in vitro and in vivo.
Two central cavity mutations severely impair function
Previous studies had implicated the central cavity as essential for Hfq and Sm protein function. Only two cavity mutants, Hfq Y55A and Hfq K56A, showed consistent in vitro defects in binding to DsrA. Both mutations target highly conserved residues within the Sm2 motif. Mutations at other conserved Sm2 residues (His57, Ile59 and Ser60) did not significantly impair in vitro or in vivo behavior.
The minimal substrate RNA-U showed much greater sensitivity to Sm2 region mutations, whereas rpoS mRNA 5' UTR and A27 were unaffected. The known
RNA-binding cavity along the inner rim of the Hfq hexamer thus seems only to
56 interact with the (A+U)-rich elements of the ncRNAs, and does not represent the
primary binding surface for poly(A) sequences or mRNAs. Additional RNA-
binding surfaces must be present on Hfq, supporting the idea that Hfq functions
by colocalizing ncRNAs and their mRNA targets. Hfq probably facilitates the
strand exchange reaction largely by putting the RNAs in close proximity, and possibly by presenting appropriate interaction surfaces toward one another.
The DsrA half-life experiments provide an additional window into the complexities of the Hfq system. Previous work has shown that in the absence of Hfq DsrA stability is markedly decreased in vivo 7. This effect is thought to be due to the
overlap of the Hfq-binding and RNase E cleavage sites on DsrA. Although both
Hfq Y55A and Hfq K56A are inactive for DsrA-mediated regulation of rpoS, their
abilities to stabilize DsrA in vivo are diametrically opposed. Hfq Y55A behaves
like wild-type Hfq whereas Hfq K56A provides DsrA almost no protection from
RNase E. The notable ability of Hfq Y55A to stabilize DsrA suggests that,
although DsrA binds to Hfq Y55A in vivo, it cannot correctly interact with rpoS
mRNA. This difference between Hfq Y55A and Hfq K56A could relate to the
stability of a ternary complex involving Hfq, DsrA and rpoS mRNA (P.J.M., E.
Espinosa, Indiana University, and A.L.F., unpublished data).
Proximal face mutations show few specific effects
Most of the proximal face mutations showed little effect on in vitro and in vivo
activity. Where disagreement was observed (Hfq D40A and Hfq F42A), the data
57 are rectified by protein accumulation defects in vivo. Although it is unclear why
these two mutants fail to accumulate, the result clearly explains their inability to
complement the hfq- strain.
Distal face mutations alter binding only to poly(A)
Distal face mutants were generated to test the hypothesis that RNA- or protein-
binding interactions might occur on that face5, 37. Hfq Y25D and Hfq I30D bound
A27 with five- to ten-fold reduced affinity, but did not show altered binding to any of the other three RNA substrates.
The distal face mutants complemented the rpoS activation function of wild-type
Hfq in vivo, and effectively bound DsrA and rpoS mRNA 5' UTR in vitro. Hfq is known to be involved in modulating polyadenylation, however. Because all mutations that altered poly(A) binding localized to the distal face, one might infer that polyadenylation control uses this surface of the protein. Further studies are required to find whether these mutations differentially affect polyadenylation.
Archaeal Sm proteins are believed to aggregate into long rodlike structures46.
Even in the absence of extended rods, a dodecameric structure could have explained well how Hfq hexamers bring together RNAs in a pairwise fashion.
Such a model requires the distal face to be involved in an Hfq-Hfq contact. The addition of six aspartates in lieu of hydrophobic residues at this potential interaction surface should have been quite destabilizing were such face-to-face
58 dodecamerization important. Our results do not absolutely preclude the formation of such species, but they do argue against it.
rpoS mRNA 5'-UTR binding remains mysterious
An unresolved issue from this work is where rpoS mRNA binds Hfq. The work of
Lease and Woodson indicates that several U-rich sequences within the rpoS mRNA 5' UTR become protected from nuclease digestion upon Hfq binding (R.
Lease and S. Woodson, Johns Hopkins University, personal communication).
Such results imply an interaction with the central cavity, but our data suggest that, if the mRNA binds the central cavity, additional contacts must mask effects of the Y55A and K56A mutations. In addition to the 5' UTR used in these studies, rpoS mRNA in vivo would contain the coding region and the 3' UTR. Does the mRNA naturally contact both faces of Hfq? If so, additional binding determinants might be present along the exterior edge of the torus. Our current study has not probed those regions. Furthermore, the C-terminal extension of Hfq could contribute to RNA binding in ways we have not yet recognized.
Implications for understanding Sm and Lsm proteins
How do these data reflect on our understanding of the eukaryotic Sm and Lsm proteins, with which Hfq shares a common ancestor? In the Pyrococcus abyssi
Sm1 core complex, RNA was shown to bind facially, near the end of strand 2
(ref. 47), and binding was significantly impaired in a Y34V mutant. This binding surface corresponds to the L3 region in the Sm1 motif of Hfq, adjacent to Asp40,
59 Gln41 and Phe42. Although the mutations at these sites do not exhibit the
marked RNA-binding defect seen in the P. abyssi Sm1 system, the accumulation defect of Hfq D40A implies that this region along the proximal face of the torus is still critical for function of the resulting RNP complex. The eukaryotic SmG protein uses this L3 region to contact RNAs in spliceosomal small nuclear ribonucleoprotein (snRNP) cores48, 49. Our data are consistent with a second site
of contact between the RNAs and Hfq, but the exact location of the L3 contacts may have evolved after the homohexameric Hfq complex diverged to form the heteromeric aggregates observed in modern spliceosomes. Current models of
U1 snRNPs also imply significant contact between the duplex regions of the RNA and the outer rim of the torus. Further exploration will probe these surfaces.
60 SUMMARY
These experiments reveal that Hfq contains at least two distinct RNA-binding surfaces. Mutations that alter ncRNA binding affinity cluster around the Sm2 domain, but do not significantly perturb affinity for rpoS mRNA 5' UTR.
Additionally, distal face mutations are the only ones tested that affect A27 binding.
Poly(A) RNA binding therefore uses contacts on the back face of the torus. Such an organization could spatially and functionally separate Hfq's effects on polyadenylation from those mediating base-pairing between ncRNAs and their mRNA targets.
61 METHODS
Plasmid construction for rpoS mRNA 5' UTR.
The 5' UTR from E. coli rpoS (nucleotides -134 to +3) was obtained by PCR using the primers rpoSA1 and rpoSA2 (Supplementary Table 2 online) using Pfu
Turbo (Stratagene). The PCR product was ligated into pUC19 using BamH1 and
EcoRI. The resulting DNA was transformed into XL-10 Gold cells (Stratagene).
Plasmid pJEF-10301 was isolated by miniprep (Qiagen) and sequenced. Large- scale isolation of plasmid DNA was done using the Qiagen Gigaprep protocol, and the vector was prepared for runoff transcription by exhaustive digestion with
SspI.
RNA preparation for in vitro binding.
RNA-U, A18 and A27 were purchased from Dharmacon Research and deprotected
following the manufacturer's protocol. RNA quality was assessed by denaturing
PAGE and gel-purified as necessary. Other RNAs (DsrA and rpoS mRNA 5'
UTR) were transcribed in vitro and gel-purified as described37.
Site-directed mutagenesis.
Using the QuikChange procedure (Stratagene), all mutants were prepared in two
separate backgrounds. The pET-21b background has a C-terminal His6-tag and
was used for in vitro analysis. The pBAD24 background is under an arabinose
promoter system and was used for in vivo analysis. All mutations were verified by
62 sequencing.
Hfq expression and purification.
Wild-type Hfq is heat-stable and the standard purification protocol exploits this
property25, 50. As several of the mutant proteins were found to be heat-sensitive,
all mutants (as well as wild-type Hfq) were purified by Co2+-affinity
chromatography, using a C-terminal His6-tag. Expression was induced by
addition of 1 mM IPTG to cultures grown to A 600 = 0.4−0.6. Induction proceeded
for 4 h at 37 °C before harvesting. Cell pellets (0.5-l equivalents) were
resuspended in 25 ml lysis buffer (50 mM HEPES, pH 7.5, 500 mM NH4Cl, 20
mM imidazole, 5% (w/v) glycerol) with EDTA-free Complete protease inhibitor
cocktail (Stratagene) and lysed by ultrasonication. Lysate was treated with
DNase I (100 U) and RNAse A (100 g) and incubated on ice for 1 h.
Centrifugally clarified lysate was passed over a Hi-trap metal chelation column
(Amersham-Pharmacia) preloaded with CoSO4. The column was washed with
five volumes of lysis buffer, then five volumes of wash buffer 1 (50 mM HEPES,
pH 7.5,1 M NH4Cl, 5% (w/v) glycerol). Hfq was eluted with five volumes of elution
buffer 1 (50 mM HEPES, pH 7.5, 500 mM NH4Cl, 250 mM imidazole, 5% (w/v)
glycerol) followed by five volumes of elution buffer 2 (50 mM HEPES, pH 7.5, 8 M
urea, 1 M NH4Cl, 50 mM EDTA, 5% (w/v) glycerol). Protein was concentrated to
0.5−1.0 mg ml-1 and dialyzed against storage buffer (50 mM HEPES, pH 7.5, 250
mM NH4Cl, 1 mM EDTA, 10% (w/v) glycerol). Concentrations of protein and any residual RNA were assessed by the Warburg-Christian method51. Purity was
assessed by SDS-PAGE. Mutants were checked by CD spectroscopy using a
63 Jasco-J715 spectrometer and by dynamic light scattering using a Malvern Nano-
S Zetasizer to ensure proper folding and oligomerization relative to wild-type Hfq.
Electrophoretic mobility shift assays.
5' end−labeled RNAs were annealed at 90 °C for 120 s, cooled to 37 °C, and
incubated with Hfq for 30 min in 50 mM HEPES, pH 7.5, 250 mM NH4Cl.
Immediately before loading, samples were diluted with an equal volume of native
loading buffer (10% (w/v) sucrose, xylene cyanol, bromophenol blue) or
denaturing loading buffer (7 M urea, 1 TBE, xylene cyanol, bromophenol blue).
Samples (15 l) were resolved at 5 W on either 5% (w/v) polyacrylamide native
gels or 5% (w/v) polyacrylamide/7 M urea gels. Dried gels were visualized by
phosphorimaging (Molecular Dynamics) using a Typhoon 9210 imaging system
(Amersham-Pharmacia). Quantification was done using ImageQuant 5.2
(Molecular Dynamics) and Kaleidagraph 3.0 (Synergy). Data were fit using
nonlinear least-squares analysis to a cooperative binding model. Cooperativity
values (n) tended to fall between 2 and 3. Higher n-values yielded comparable
goodness-of-fit parameters. In the case of the A27 competition assays, complexes
of DsrA or rpoS mRNA 5' UTR with Hfq were preassembled.
Isothermal titration calorimetry.
Calorimetry was done on a MicroCal VP-ITC. Samples were dialyzed into
reaction buffer (50 mM HEPES, pH 7.5, 250 mM NH4Cl, 10% (w/v) glycerol) and
degassed before loading each experiment. RNAs (21−25 M) were titrated into
64 1.4 ml of 0.6−3.5 M Hfq hexamer over 35 8- l injections with constant stirring at
310 r.p.m., 4-min injection spacings, and thermostatting at 25 °C. Data were
corrected by subtraction of a baseline defined by the terminal 10−15 injection points after saturation of the binding event. Data were analyzed using Origin 7.0
(MicroCal) as described52.
Western blot analysis.
Total cellular extracts of E. coli were separated on tris-tricine SDS 16% (w/v) polyacrylamide gels53 and electroblotted54 to PVDF membrane. Equal loading
across lanes was verified by staining with Ponceau S (Sigma). The membrane
was probed with rabbit anti-Hfq polyclonal antisera7. Antibody−antigen complex was visualized with goat anti-rabbit immunoglobin horseradish-peroxidase- conjugated antibody (Pierce) and ECL reagent kit (Pierce).
Purification of RNA for half-life studies.
DsrA half-life was determined as described37, 55. Cultures of wild-type and hfq-
-1 strains were grown at 30 °C to A 600 = 0.4−0.6. Rifampicin (100 g l final
concentration) was added and cells were collected at the indicated time points.
These samples were added immediately to prechilled tubes containing 0.1
volume phenol stop solution (5% (v/v) buffered phenol, pH 7.4, in ethanol)56.
Cells were pelleted, resuspended in 200 l TE (10 mM Tris, pH 8.0, 1 mM
EDTA), and passed over QIAshredder mini columns (Qiagen). RNA was
extracted with Trizol reagent (Invitrogen), precipitated with isopropoanol, and
65 washed with 75% (v/v) ethanol. RNA was incubated with 30 U of RQ1 DNase
(Promega) at 37 °C for 20 min followed by DNase inactivation at 60 °C for 10
min. Trizol extraction and isopropanol precipitation were repeated before
resuspension of RNA in RNase-free water. Concentrations were determined by
measuring absorbance at 260 nm.
Quantitative RT-PCR (QRT-PCR).
Relative RNA concentrations were determined by quantitative RT-PCR using a
Roche LightCycler TaqMan PCR system (Applied Biosystems). Primer and probe
design was based on the E. coli dsrA and 5S RNAs using Primer Express 1.5
(ABI). The dsrA primers and probes were designed to detect only full length
DsrA. Primers and probes were chemically synthesized (IDT). Probes were 5'
end−labeled with 6-carboxyfluorescein and 3' end−labeled with 'black hole quencher' BHQ1 (Biosearch Technologies). Each 10 l RT-PCR mixture
contained 50 ng RNA, LightCycler-RT-PCR reaction mix hybridization probe
(Roche), LightCycler RT-PCR enzyme mix (omitted in RT-negative reactions),
300 nM each of the forward and reverse primers, and 250 nM Taqman probe.
Both RT-positive and RT-negative reactions were tested in duplicate. Reactions
were performed using a Roche LightCycler with optimized cycle conditions. DsrA
RNA abundance relative to 5S ribosomal RNA was determined using the - Ct method as described by the manufacturer (ABI). 5S RNA was used as the internal control.
66 In vivo RpoS::LacZ fusion activity.
RpoS::LacZ fusion activity was determined as described7. Briefly, cells were
grown at 30 °C in LB medium with the appropriate antibiotic. Total -
galactosidase units were determined as described57 and plotted against the
culture A 600. The slope of the linear regression (differential rate of expression)
was used as the specific activity. Mutant activities were determined at least three
times and s.d. calculated.
Note: Supplementary information is available on the Nature Structural &
Molecular Biology website.
67 Figure 1. Sequence alignments and structure of Hfq.
(a) Sequences 1412.11 and 1923.1 are entries from the conserved domain database representing the superfamily of Lsm proteins and Hfq, respectively.
The two conserved domains have been aligned to properly juxtapose the motifs present in both protein families. Sequence elements in red represent the most highly conserved portions of the domain. HfqEC is the E. coli Hfq protein, colored based on an alignment of 25 bacterial Hfq homologs showing complete identity and sequence similarity in red and green, respectively. Above the alignment, triangles mark sites where RNA is known to interact with Lsm proteins and diamonds indicate known RNA-binding interactions with Hfq. The bottom line of the alignment shows the sites mutated as part of the study; P, location on the proximal face; D, location on the distal face; C, central cavity mutation.
Numbering is provided for reference and is based on the E. coli Hfq sequence.
Secondary structure information derives from the crystal structure of HfqEC (PDB entry 1HK9)27. (b) Space-filling representation of the HfqEC crystal structure
(PDB entry 1HK9) color-coded with the sequence conservation data from the
HfqEC alignment in a. The distal face representation shows the same image rotated 180° relative to the proximal face around a vertical axis in the plane of the page. (c) Space-filling representation of the HfqEC crystal structure upon which the composite RNA binding data from Table 1 have been superimposed.
68
69 Figure 2. In vitro analysis of RNA binding to mutant Hfq proteins.
(a) Gel shift experiments showing the binding of wild-type Hfq to DsrA, RNA-U,
rpoS mRNA 5' UTR and A27.
(b) Quantification of the gel shift experiments in a. Closed circles, DsrA K1; open
circles, RNA-U; closed squares, rpoS mRNA 5' UTR; open squares, A27. Lines
represent nonlinear least-squares fitting to a cooperative binding model.
(c) Histogram showing the free energy of binding relative to wild-type (WT) Hfq for each Hfq mutant. Gs between -0.5 and 0.5 kcal mol-1 indicate insignificant effects on RNA binding. Asterisks indicate data that are lower limits of the actual
effect as the interactions were too weak to be measured. Blue, DsrA−Hfq6; red,
DsrA−(Hfq6)2; green, RNA-U; brown, rpoS mRNA 5' UTR; lavender, A27.
70
71
72 Figure 3. Native gel analysis showing the effect of A27 addition to the
DsrA−(Hfq6)2 complex.
DsrA−(Hfq6)2 was preassembled at appropriate concentrations of Hfq (based on
the measured affinity constants in Table 2) using 32P-labeled DsrA before the
addition of unlabeled A27 (0, 0.03, 0.1, 0.3, 1 or 3 M). Gels compare the
behavior of wild-type (WT) Hfq to Hfq Y25D and Hfq K56A as labeled. A
supershifted species is observed (arrow) only in the case of WT Hfq. For Hfq
Y25D, the addition of A27 had no effect on DsrA binding. For Hfq K56A, A27 displaced DsrA in the form of the homodimer, DsrA2, based on its gel migration
against authentic standards.
73
74 Figure 4. Assay for in vivo accumulation and rpoS activation.
(a) Schematic diagram of the reporter construct used for the Hfq
complementation assay.
(b) Western blot showing the accumulation of Hfq in vivo. In the control lanes, + and - refer to the presence or absence of 150 M arabinose, added to induce Hfq
from an araBAD promoter. Vector represents the pBAD plasmid45 without the hfq gene. The Hfq lane shows the endogenous Hfq levels in an hfq+ strain of E. coli.
(c) Histogram showing the results of in vivo translational activation of the
RpoS::LacZ fusion protein for wild-type (WT) and mutant Hfq proteins. Values <4
are considered significant defects based on the error in this assay. -gal, - galactosidase activity.
75
76 Figure 5. Lifetime analysis of DsrA in vivo.
Data are based on QRT-PCR for wild-type (WT) Hfq and selected mutants using
primers specific for full-length DsrA. Solid lines (Hfq mutants) and dashed lines
(Hfq and hfq-) are least-squares fits to the data from which the half-lives were
calculated. Hfq K56A has been fit to a double-exponential decay whereas the other data are fit to a single-exponential model.
77
78
Table 1 Comparison of in vitro Affinity Data with in vivo Activities for Hfq Mutants.
aDsrA exhibits two sequential transitions – a shift and a supershift. The first value
corresponds to K1 and the second value corresponds to the supershift, K2. Close inspection of the rpoS mRNA 5’-UTR shifts reveals two apparent shifted species similar to those seen with DsrA. However, the two transitions are sufficiently coupled that they cannot be fit separately.
79 bHfq- strains have a DsrA half-life of 5 ± 2 min.
cThese data display a biphasic time dependence 90% of the DsrA RNA decays with the half-life of 0.9 min. The final 10% of the RNA displays a much longer half-life of approximately 35 min.
d No clear distinction between K1 and K2 behavior was possible due to abnormal
smearing of the bands. Data were fit to a single transition.
80
Table 2. Summary of A27 Competition Experiments.
Complex Mutant [Hfq] Outcome upon A27 (nM hexamer) Competition
DsrA•Hfq6
WT 42 No effect
Y25D 25 No effect
K56A 125 Abnormal
competition
DsrA•(Hfq6)2
WT 125 Supershift
Y25D 83 No effect
K56A 250 Abnormal
competition
rpoS•(Hfq6)2
WT 83 Supershift
Y25D 50 No effect
K56A 125 No effect
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89 MANUSCRIPT II
Translational Coupling of the Endonuclease Gene to That of its Upstream
Regultor in thr PvuII type II Restriction-Modification System
(Meenakshi Kaul Kaw and Dr. R. M. Blumenthal
90 ABSTRACT
Type II restriction-modification (RM) systems have two independent catalytic components: a modification methyltransferase which methylates and thus protects the DNA, and a restriction endonuclease. Transcriptional mechanisms that delay endonuclease expression have already been identified. A substantial subset of RM systems is controlled by an unusual family of small, broad host-range transcription activators called C proteins. In the PvuII sytem from Proteus vulgaris, C.PvuII activates its own gene’s transcription, along with that of the downstream endonuclease gene. In theory this acts like a timed switch, giving low R.PvuII levels early but then rapidly ramping them up. Studies of C-controlled type II RM system regulation have focused on transcriptional level. In all these systems there is a ±20 nt overlap between the C termination codon and the R (endonuclease) initiation codon. This consistent overlap suggests possible translational coupling. In addition pvuIIR is preceded by hairpins that might regulate translation by sequestering the ribosome binding site.
Analysis of reporter fusion strains, together with real time (RT-PCR) measurements of mRNA levels indicate that the putative hairpins play no detectible role in regulating the PvuII RM system, but that the translation of pvuIIR is coupled to that of pvuIIC.
91 INTRODUCTION
Bacterial type II restriction-modification (R-M) systems are abundant in
both the prokaryotic and the archaebacterial worlds (Kessler and Manta, 1990).
Some of these are specified by plasmids, while others are chromosomal. A type
II RM system is composed of genes that encode a methyltransferase (MTase) and a restriction endonuclease (REase). Many type II RM systems are genetically mobile and shift from one cell to the other by transduction, transformation or conjugation (Vijesurier et al., 2000; Naderer et al., 2002;
Knowle, et al., 2005).
PvuII is a type II RM system from the Gram-negative bacterium Proteus vulgaris (Gingeras et al., 1981). The DNA methyltransferase (M. PvuII) modifies the recognized DNA sequence CAGCTG by methylation of the internal cytosine residue, creating N4-methylcytosine (Blumenthal et al., 1985) and the restriction endonuclease (R. PvuII) cleaves the central GpC of the same unmethylated sequence (Gingeras et al., 1981). Methylation protects the host DNA from being cleaved by REase. This endonuclease is allowed to cleave the incoming unmethylated foreign DNA, acting as primitive immune system, especially against the bacteriophages (Raleigh and Brooks, 1998). However, both components act independently of each other. Strict regulation is needed to properly time the expression of the two components of the RM system while
92 establishing themselves in the new host, so as to delay REase until MTase
modifies the host DNA (Vijesurier et al., 2000).
The mechanism behind this regulation was explored by our laboratory and
other research groups. In 1991 Brooks and her colleagues, and Tao and
colleagues, from our lab discovered a third ORF which was designated as C
(controller) protein in the BamHI and PvuII systems respectively. Based on
sequence comparison, similar ORFs were identified in the SmaI and EcorV
systems as well (Tao et al., 1991). Now this C protein has been found in BglII,
Esp1396I, Eco72I, AhdI, Kpn2I, EcorV, EcoO109I, SmaI, SgrAI, LIaDI, SptAI,
BcnI, NmeSI, BatLVI, MunI, SnaBI and couple more are being characterized.
A conserved sequence was detected upstream of the C gene, known as
“C- box” and was suggested to be a C protein binding site. C.PvuII binds the C-
box and induces transcription of the PvuII genes (Vijesurier et al., 2000). C.PvuII
regulates its own transcription as well as that of the of downstream REase gene,
which lacks its own promoter. (Vijesurier et al., 2000; Knowle et al., 2005).
PvuII.C associated transcriptional mechanisms that delay endonuclease
expression have already been identified by our lab (Tao et al., 1991; Tao et al.,
1992). Such mechanisms are regulated by two promoters, one that is weak but independent of the C. PvuII, and the other promoter is strong but dependent on
C. PvuII. It was suggested that transcript initiation within the C boxes is the initial,
93 C-independent transcription of pvuIICR, while the C-dependent transcripts are
leaderless mRNA present further downstream at the initiation codon of the pvuIIC
gene (Knowle, et al., 2005). The importance of this delay is highlighted by the
fact that pre-expressing C protein prevents introduction of an intact RM system
(Vijesurier et al., 2000; Kobayashi, 2001).
It is not clear if this transcription-level regulation is sufficient to protect the cell. The early transcripts from the weak C-independent promoter should still go into pvuIIR. We accordingly investigated whether there is some type of translational control. There is a pvuIIC – pvuIIR overlap, and pvuIIR is preceded by potential hairpins, in both cases suggesting possible translational control of pvuIIR. C.PvuII could in theory perform additional control. Figure 1 illustrates a
±20 nt overlap region between the C terminator and R (endonuclease) initiator in
a selection of C-dependent RM systems. We investigated whether this consistent
overlap implies translational coupling or translational regulation.
94 MATERIALS AND METHODS
Cloning and generation of mutants.
The Cla and Esp mutants of pvuIIC were generated by Tao et al (1991).
pPvuRM3.4 was generated by cloning the PvuII RMS2 as a 3.4 kb fragment into
pBR322. The pPvuRM3 includes two unique enzyme sites in the pvuIIC ORF,
ClaI and EspI sites and were used to generate the mutants. pvuIIC includes two
unique sites: ClaI and EspI (BlpI). At the ClaI site two frameshift mutations were generated either by filling in the 5’ extensions with Klenow polymerase or
digestion using mung bean nuclease. EspI yields 3-bp overhang (GC/TNAGC),
so that Klenow fill-in or digestion with mung bean nuclease created an in-frame
mutation by deletion or insertion of a leucine codon. These mutants were
sequence confirmed.
PCR amplification.
EspI mutants (Esp19 with insertion and Esp33 with deletion of leucine
codon) and Cla mutant (Cla35 insertion mutation) derivatives of pPvuII RM3.4
along with WT were purified via QIAprep plasmid DNA purification kit (Qiagen).
Two primers were purchased from Integrated DNA technology company (IDT).
An XmnI site was added on the 5’ end of the insert and an EcoRI site was added
to the 3’ end of the insert.The primer set was used to amplify the fragment
between XmnI that included ATG of pvuIIC and few codons into pvuIIR including
the overlap between pvuIIC and pvuIIR (~285 bp). PCR amplification used high
95 fidelity Pfx polymerase enzyme (New England Biolabs) was used to amplify the
fragment. The XmnI and EcoRI digested fragment was purified by the Qiagen gel
extraction kit. A similar protocol was followed for amplification as well as
purification for all three mutants including the wild type.
Ligation and transformation and sequence confirmation.
The amplified fragments were cloned into TOPO 2.1 vector from TOPO
TA cloning kit (Invitrogen). The ligation products were transformed into TOP-10
chemically competent cells (Invitrogen). Transformation products were plated
onto LB agar with ampicillin 100 µg/ml. Transformants were inoculated into LB
broth with appropriate antibiotics. Clones that demonstrated the expected pattern
on gel electrophoresis analysis were confirmed by sequencing.
Construction of LacZ translational fusions.
These inserts from sequence confirmed clones in TOPO 2.1 were again
amplified using a second set of primers purchased from Integrated DNA technology company (IDT). The primer set was used to amplify the fragment between ‘TG’ of the ATG of pvuIIC and few codon into pvuIIR. EcorI was added to the 3’ end of the insert. PCR amplification used high fidelity Pfx polymerase
(New England Biolabs). The EcoRI digested fragment was purified by electrophoresis and the Qiagen gel extraction kit. A similar protocol was followed for all three mutants and the wild type.
96 Ligation and transformation and sequence confirmation of the translational
fusions.
We constructed translational fusions with both C and R being in-frame
alternatively with the lacZ gene of plex3b (ATCC# 87200). For the mutants to have pvuIIC in-frame with lacZ, a 30bp oligonucleotide was acquired with a BglII site at the 5’ end (sense strand) and EcoRI at the 3’ end. In the middle we inserted a unique XbaI site that facilitated identification of the desired clones.
This DNA was cloned between the XmnI and EcoRI site of the pLEX3B vector, in such that ‘A’ of the XmnI site blunt ligated to the ‘TG’ of the insert on the 5’ end;
at the 3’ end ligation used the EcoRI site.
Ligation products were transformed into TOP10 chemically competent
cells (Invitrogen). The transformation product was plated onto the LB agar with
ampicillin (100mg/ml), IPTG (1 mM), and X-gal. Transformants that turned blue
were patched and also inoculated into LB broth with the appropriate antibiotics.
Putative lacZ translational fusions were isolated and confirmed by restriction-
digestion as well as colony PCR. Clones that demonstrated the desired pattern
on gel electrophoresis analysis were confirmed by sequencing.
Site-directed mutagenesis.
In order to bring all these mutants as well as the WT with pvuIIR in-frame
with lacZ gene of the pLEX3B, we deleted two bp using site-directed
mutagenesis. The primers (IDT) were designed to function with the QuickII site-
97 directed mutagenesis protocol and the kit from Stratagene. We had all three mutants and wild type (WT) both with pvuIIC and pvuIIR in frame with lacZ gene.
While changing the C translational fusions into R translational fusions, the stop
codon TAA was shifted, so we restored the stop codon to its original position by using Quikchange II protocol. Next these mutants were provided with a wild type pvuIIC (C+) and mutant (C-) in trans from a second compatible plasmid.
Site-directed mutagenesis for deletion of the hairpin.
We also made an in-frame arm1 and arm 2 deletions in the predicted hairpin using site-directed mutagenesis kit and protocols. Again this set was also confirmed by sequencing (MWG). The hairpins are described in detail in the results section.
The wild type (WT) pvuIIC translational fusion contains the full C ORF and few codons from the pvuIIR and is in-frame with LacZ reporter gene of the pLEX3B. We had to remove the stop codon from the C ORF fuse with LacZ. In the deletion derivative of this we have three mutations consisting of Cla35 which is out of frame with its natural reading frame in the native context. However, we have got this in-frame with the LacZ reporter gene to create a translational fusion by deleting the region between BglII and EcoRI region and filling in with 30bp oligonucleotide along with a unique XbaI site. We used a similar technique to create translational fusions of other two mutants including Esp19 and Esp33. The two (AG) nucleotides were deleted to shift the frame, such that wt pvuIIR along
98 with the mutant pvuIIR would be in-frame with the lacZ gene of the plasmid pLEX3B. The Shine dalgarno (SD) sequence and the promoter from the vector were included in the fusions. This shifted the stop codon for pvuIIC by few codons. We used site directed mutagenesis to restore the stop codon to its original position by substitution. The Esp33C fusion was created by insertion of
30 mer oligonucleotide. The pvuIIR has its own translation initiation region and also a SD sequence. We examined all these fusion constructs by Beta- galactosidase assay.
Beta-galactosidase assay.
Beta-galactosidase assay was used to analyze the expression of
translational fusions. We measured the hydrolysis of the substrate ο -
nitrophenyl-β-D-galactoside (ONPG) that is colorless and yields a colored
product on hydrolysis into galactose and ο - nitrophenol by the enzyme β
galactosidase.
Overnight cultures were grown at 37 ο C, in the presence of tetracycline
antibiotic (10 µg/ml) and IPTG (1 mM) for induction. This culture was diluted
1:100 in MOPS rich defined medium, and the cultures were grown to exponential
phase. Samples were collected at regular intervals and the optical density (OD)
at 600 nm was measured for every 1 ml sample. Two drops of chloroform and
one drop of 0.1% SDS were added, and the sample vortexed for about 10
seconds to open the cells. Samples were placed in a 28ο C water bath for 5 min, 99 and 200 µl ONPG (4 mg/ml) are added to each tube. The time of ONPG addition
of and the time when the color changed to yellow were noted. To stop the
reaction, 500µl Na2CO3 (1 M) were added.
For each sample we measured the OD at 420 nm. Using the formula of
Miller (1972) we calculated the units of beta-galactosidase. Miller units were
plotted against the culture OD 600 nm. The slope from linear regression
(differential rate of expression) indicated specific activity. Some of the constructs were assayed in triplicates and standard error was calculated (Miller, 1972).
Growth experiment and sample isolation.
Overnight culures were grown in MOPS rich medium, with ampicillin as an
antibiotic (100µg/ml). Next day the samples were diluted 1/100 and allowed to
grow to an OD 600nm (of ~0.3) along with IPTG (1mM). Samples for real time RT-
PCR analysis were isolated at the indicated OD from the flask and added to the
two volumes of RNA stabilization buffer (RNA Protect Bacteria Reagent, Qiagen,
Valencia, CA). This inhibits the changes that would occur in the mRNA content if
bacteria are harvested. Samples were mixed at left at room temperature for
10min.
RNA isolation and cDNA synthesis.
RNeasy miniprep kit (Qiagen) was used for isolating the total RNA. Cells
in stabilization buffer were harvested by centrifuging at 4ο C for 15 min at 5,000 100 rpm. Pellet was resuspended in 1x TE buffer containing lysozyme (400µg/ml)
after removing the supernatant. Ethyl alcohol was added to precipitate the RNA,
and then the coumns were washed and eluted. RNA was treated with RQ1
RNAse-free DNAse (Promega, Madison, WI) to remove DNA as per their
directions. CDNA was synthesized using toptal RNA as template, random
hexamers (Invitrogen, Calsbad, CA), and ImPromII reverse transcriptase
(Promega). The random primers were annealed at 25ο C for 5 min, and the first
strand was then extended at 42ο C for 1 hour. The reverse transcriptase was then
inactivated by heating at 70ο C for 1 hour.
Real time RT-PCR analysis.
A primer set was designed for the region including pvuIICR genes from the IDT (Coralville, IA). Before the experiment dilutions of cDNA were tested to
determine the maximal efficacy concentration desired for amplification, and also
the efficiency for each primer. A house keeping gene recA expression was also
measured and the expression levels of WT, Cla 35, Esp19 and Esp33 mutant
were normalized to the RecA levels. Cycle threshold (CT) values were detected
by Roche Light Cycler using SYBR GREEN fluroscence method. Melting curve
was used to confirm the formation of specific RNA. The standard curve method was used to determine the relative amounts of mRNA which were normalized to
recA (Wong and Medrano, 2005).
101 RESULTS AND DISCUSSION
Plasmid pPvuIIRM3.4 carries the WT PvuII RM system in pBR322
(Blumenthal et al., 1985). The pvuIIC gene contains two unique sites (ClaI and
EspI) that were used to generate mutants (Tao et al., 1991). Wild type and
mutant versions of pPvuIIRM3.4 were purified and used as a template to amplify
the region that included TG of the pvuIIC initiator and some codons from pvuIIR;
this was cloned to generate lacZ translational fusions in pLEX3B (Diederich et al., 1994).
Previous work from our laboratory demonstrated a sharp decrease in the expression of pvuIIR in four pvuIIC mutants (Tao, et al., 1991). The mutants showed greatly-impaired restriction of unmethylated bacteriophage λ or plasmid pACYC184 and the in vitro REase activity was 104-fold less than in strains that
carried the intact parental plasmid. Providing pvuIIC in trans had no effect on the
wild type strain, but there was a 104 fold increase of in vitro PvuII REase activity
and the restriction of λvir increased by 107 fold.
The mutants tested included two in-frame (EspI) and one frameshift (ClaI)
mutants. These results reveal that pvuIIC complements both in frame as well as
out of frame mutants, reducing the likelihood of strong translational coupling.
However in that study the measure of pvuIIR expression was restriction of
bacteriophage lambda, which is not necessarily a linear and quantitative 102 measure of pvuIIR translation. The lacZ fusion measurements would more accurately address this question.
LacZ measurements are made over the course of a growth experiment.
The plot of activity vs. culture density is linear if the measurements of cells are in balanced growth. The resulting slopes give a precise measure of specific activity.
Effect of C.PvuII on the expression of its own gene (pvuIIC) and that of the restriction endonuclease (pvuIIR).
In majority of C - associated RM systems that have been characterized to date, the gene for the regulatory protein precedes the REase gene, suggesting cotranscription and coregulation at the transcriptional level. These predictions have been confirmed for the PvuII system (Vijesurier et al 2000). Bacterial genes are often cotranscribed in a polycistronic mRNA. Many operons require translation of preceding gene for the efficient translation of subsequent gene.
This phenomenon is called translational coupling, and was observed for the first time in the tryptophan operon of E. coli (Oppenheim and Yanofsky, 1980).
Studies on gene regulation of another RM system Esp1396I
(Cesnaviciene, et al., 2003) demonstrated that C.Esp1396I activates the expression of its own gene and downstream gene for REase and that the R and
C genes exhibit coupled translation. Another study on Eco72I (Rimseliene, at al.,
1995) revealed that providing C.Eco72I in trans to lacZ translational fusion
103 plasmids of eco72IR::lacZ restored the REase activity when eco72I gene was inactivated by the frameshift mutation. These findings suggest that C.Eco72I translationally regulates the REase gene expression.
In order to test for coupling of pvuIIC and pvuIIR, we measured β-gal activity from WT fusions. Specific activity was measured with the pvuIIC stop
codon moved ~ 20 nt downstream or at its native site (Fig 4). The level of
expression of WT R-translational fusion was not significantly different from that of the C-translational fusion. Providing WT C.PvuII (or a mutant version) in trans from another, compatible plasmid had no effect on the fusions (Fig 6). The native promoter was replaced by Plac in these plasmids, so C.PvuII should have no
transcriptional effects.
Based on previous findings (Tao et al., 1992) and current results from the
WT translational fusions, the expression of pvuIIR is comparable and in proportion with that of pvuIIC suggetsing that they are translated at the same rate.
Frame shift mutation in pvuIIC affects translation of pvuIIC and pvuIIR.
In most polycictronic operons the termination codon is located very close
to the initiation codon of the downstream gene; in some cases they even overlap
(Oppenheim and Yanofsky, 1980). Such overlaps may result in well coordinated
expression of the genes, with the translational efficiency of the downstream gene
104 often dependent on expression of the preceding gene. For example, the
galactose operon exhibits translational coupling, with the expected effects of
frameshift and nonsense mutations (Shumperli, et al., 1982).
To test whether pvuIIR translation is coupled to that of pvuIIC, we investigated the Cla35 mutant, which is a frameshift mutation in pvuIIC that
introduces a terminator 80 nucleotides upstream of the normal stop codon (Tao
et al., 1991; Fig 2). lacZ fusions in the pvuIIR translational frame showed reduced
levels of expression (Fig 7). The same mutation with lacZ fused in pvuIIC frame
shows a drop by the same factor. There was no difference between the native
and downstream (shifted) stop codon for C (see Fig 4). Providing WT (or mutant)
pvuIIC in trans did not restore expression. This parallel drop is consistent with
coupling. However, it was possible that premature translation termination led to
premature transcript termination (Shumperli, et al., 1982; Cesnaviciene, et al.,
2003). This was tested (see below).
Inframe insertion mutation does not affect expression of pvuIIC or pvuIIR.
Esp19 is an insertion mutation of one leucine codon into pvuIIC (Tao et
al., 1991) and is thus an in-frame mutation. LacZ assays performed on both C
and R translational fusions showed identical activity levels (and no change
between C+ and C- backgrounds). Since an in-frame mutation shows unchanged
expression of both pvuIIC and pvuIIR while the frameshift Cla35 mutant shows
significantly lower activity in both frames, these results support the hypothesis
105 that pvuIIR translation is coupled to that of pvuIIC. This activity is not enhanced by providing C+ from another plasmid indicating that it is entirely a cis-effect and is not dependent on the translation product of pvuIIC.
In-frame deletion mutation affects expression of pvuIIR.
Esp33 is a deletion mutation of one leucine codon in pvuIIC (Tao et al.,
1991). This mutant shows no change in the activity levels in C translational fusions. However in the R frame translational fusions, the activity is almost tripled
(in presence of C+ as well as C-). The deleted leucine codon is far from the initiator (44 codons), and presumably does not affect pvuIIC translation initiation.
Quantative RT-PCR (QRT-PCR) results confirm translation coupling phemomenon.
To test whether the apparent coupling is real, and not due to transcriptional polarity in the Cla35 mutant, relative mRNA levels were determined by quantitative RT-PCR using Roche LightCycler SYBR Green approach. The relative mRNA levels were almost equal between the WT and the mutants (Fig. 9) In fact the levels of pvuIIR mRNA in the Cla35 mutant were slightly higher, but its specific activity in beta-galactosidase assay was far lower than the WT and the in-frame Esp mutants. These findings ok the hypothesis that pvuIIR translation is coupled to that of pvuIIC.
106 Alternative hairpins demonstrated no effect on expression.
The pvuIIR gene is preceded by two alternative, predicted hairpins
(Vijesurier et al., 2000; Fig10A). These hairpins might also regulate translation of
pvuIIR because the downstream hairpin would occulude the RBS. Altering Arm 2
should disrupt both the hairpins and increase pvuIIR translation. Altering Arm1 should disrupt the 5’ hairpin (1:2), promote formation of the 3’ hairpin (2:3) and thus decrease translation of pvuIIR. To investigate whether the alternative hairpins had some role in regulation of translation we deleted Arm1 from the WT construct, the theory is that when Arm1 is deleted, Arm2 is freed to block the
RBS. This was tested for both C and R-LacZ translational fusions. The ∆ arm1 fusion expression in the pvuIIR frame is close to WT level, with a 20% difference.
The pvuIIC fusions showed no difference, as expected, since the hairpins were not believed to affect pvuIIC translation.
When Arm 2 was deleted, the theory is that the RBS would be freed and pvuIIR translation should increase. In fact, it appears that ∆ Arm 2 has no effect.
(Fig. 10B). The hairpins thus play no detectible role. It is also possible, however, that the hairpins help explain translational coupling to pvuIIC by blocking the entry of new ribosomes at the pvuIIR RBS.
107 Summary
The expression levels from assays on WT and in-frame insertion mutations
showed no difference with lacZ fusions to the pvuIIR frame or pvuIIC frame or
with or without C.PvuII being provided in trans. However, a frame shift mutation
dropped levels of expression considerably in both frames, and providing C.PvuII
in trans had no effect. These results are consistent with translational coupling.
However we wanted to exclude the possibility of transcriptional polarity from the
Cla35 mutant. Next we performed QRT-PCR using Cyber Green. As expected
the relative mRNA levels of pvuIIR Cla35 mutant were almost equal to other
variants and the WT pvuIIR lacZ fusions, though on a slightly higher side.
However, the specific activity of pvuIIR Cla35 mutant was far lower than the WT
and the pvuIIR Esp mutants. Thus, supporting the fact that the results are purely translational, and not transcriptional. Moving the pvuIIC termination codon relative to the pvuIIR initiator had no apparent effect which confirms that this
coupling is not an artifact of the shifted position of the pvuIIC stop codon.
However, the hairpins seem to play no detectible role.
108 Figure 1. Overlap regions of + 20 nucleotides between the C terminator codon and R initiator, in a selection of C- dependent RM systems
109
.
110 Figure 2. A fragment (1-280 bp) includes the region from initiator codon of pvuIIC and few codons of pvuIIR from pPvuIIRM3.4. pvuIIR has its native RBS.
Two primers used for amplification are indicated by arrows. pvuIIC includes two unique sites: ClaI and EspI (BlpI). The rest of the fragment (from 280bp downstream) shows the vector sequence and part of the lacZ gene.
111
Forward primer
ATGAGCAGAA ATACAAATCC TTTATCAGCC CGATTAACCC TTGCAAAGAA TGTAAAAAAA M S R N T N P L S A R L T L A K N V K K> pvuIIC start
70 80 90 100 110 120 ATGCGAGGCG AGCTAGGTCT ATCCCAAGAA AGCTTAGCTG ATCTAGTGGG AATCCATAGA M R G E L G L S Q E S L A D L V G I H R>
EspI site
130 140 150 160 170 180 ACCTACATTG GTTCAATTGA ACGAGCGGAA AGGAATATAT CGATAGACAA CATTGAGCGA T Y I G S I E R A E R N I S I D N I E R>
ClaI site
190 200 210 220 230 240 ATAGCAAATG CCTTAAATGT TTCTATATCA ATACTAATGA TGGAACACGA AAATGAGTCA I A N A L N V S I S I L M M E H E N E S> * Q M P * M F L Y Q Y * * W N T K M S H> Reverse primer pvuIIR start
250 260 270 280 290 300 CCCAGATCTG CAAATCTACG TCTACGCGGA ATTCCCGGGG ATCCGTCGAC CTGCAGCCAA P R S A N L R L R G I P G D P S T C S Q> P D L Q I Y V Y A E F P G I R R P A A K> Polylinker region
310 320 330 340 350 360 GCTTGCTCCC GTCGTTTTAC AACGTCGTGA CTGGGAAAAC CCTGGCGTTA CCCAACTTAA A C S R R F T T S * L G K P W R Y P T *> L A P V V L Q R R D W E N P G V T Q L pLEX3B-lacZ start C stop
370 380 390 400 410 420 TCGCCTTGCA GCACATCCCC CTTTCGCCAG CTGGCGTAAT AGCGAAGAGG CCCGCACCGA S P C S T S P F R Q L A * * R R G P H R> R L A A H P P F A S W R N S E E A R T D>
112 Figure 3. A plasmid from ATCC (#87200) was used to construct the translational
fusions; SD and Plac promoter from the vector were used to measure the beta-
galactosidase activity. The vector was digested with XmnI and EcoRI. The WT
and all three mutants were cloned between the XmnI and EcoRI site such that ‘A’ of the XmnI site blunt ligated to the ‘TG’ of the insert on the 5’ end; at the 3’ end ligation used the EcoRI site.
113
Hi ndI I I Shine-Dalgarno Xmn1 p vuIIC pvuIIR Not1 (285bp) pMB1PMB1 EcoR1 MCS
lacZ
Not1
LacZ gene
(Adapted from Diederich et al., 1994)
114 Figure 4. A 30 bp oligonucleotide was cloned between the BglII and the EcoRI site (red) to put pvuIIC in-frame with the lacZ gene of plasmid pLEX3B
(Diederich, L., 1994). A unique XbaI site was introduced in the middle to help
identify the desired clones. Site-directed mutagenesis was used for deletion of
AG (purple) in the XbaI site to bring all the mutants and the WT with pvuIIR in-
frame with the lacZ gene. A stop codon was introduced by substitution, to restore
it back to its native location (green).
115
RTIR BglII 30 MER OLIGO EcoRI GGAACACGAAAATGAGTCACCCAGATCTGCAAATCTAGACGTCTACGCGGAATTCWTC lacZ GGAACACGAAAATGAGTCACCCAGATCTGCAAATCTAGACGTCTACGCGGGTGAATTC Esp33 C lacZ GGAACACGAAAATGAGTCACCCAGATCTGCAAATCTAGACGTCTACGCGGGTGAATTC Esp19C lacZ GGAACACGAAAATGAGTCACCCAGATCTGCAAATCTAGACGTCTACGCGGGTGAATTC ClA35C lacZ
GGAACACGAAAATGAGTCACCCAGATCTGCAAATCTAGACGTCTACGCGGGTGAATTC Esp19R lacZ GGAACACGAAAATGAGTCACCCAGATCTGCAAATCTAGACGTCTACGCGGGTGAATTC Cla35R lacZ GGAACACGAAAATGAGTCACCCAGATCTGCAAATCTAGACGTCTACGCGGAATTCTGA WTR lacZ
GGAACACGAAAATGAGTCACCCAGATCTGCATAACTACGTCTACGCGGGTGAATTC Esp19R lacZ GGAACACGAAAATGAGTCACCCAGATCTGCATAACTACGTCTACGCGGGTGAATTC Cla35R lacZ GGAACACGAAAATGAGTCACCCAGATCTGCATAACTAGACGTCTACGCGGAATTC WTR lacZ GGAACACGAAAATGAGTCACCCAGATCTAAATAAATTATTAGAGGAATTCC Esp33 R lacZ
116 Figure 5. (A) Beta-galactosidase assay (WT) performed in triplicate, in C+ (y- axis) and C- (x-axis) backgrounds. Fusions were in frame with either pvuIIC
(open circles), or pvuIIR (closed circles). Specific activities were determined via linear regression plotting modified Miller units versus culture density with R2 values of 0.98 and 0.99 respectively. (B) Repeat of experiment shown in (A) but with TAA of pvuIIC restored to its native location. R2 values were 0.98 and 0.98.
117 A B
80 100
70 80 60
50 60
40
C+ LacZ 40 30
20 20
10 activity (Modofied miller units) LacZ
0 0 0 1020304050607080 0 0.2 0.4 0.6 0.8 1 1.2 C- LacZ OD 600 nm
118 Figure 6. (A) Beta-galactosidase assay (Cla 35 mutant), in C+ (y-axis) and C-
(x-axis) backgrounds. Cla35-pvuIIC (open squares), Cla35-pvuIIR (closed squares) and WT pvuIIC (open circles), and pvuIIR (closed circles) represent constructs that are in-frame with lacZ gene. Specific activities were determined via linear regression plotting modified Miller units versus culture density with R2 values of 0.98 and 0.99 respectively and compared to WT. (B) Repeat of experiment shown in (A) but with TAA of pvuIIC restored to its native location
R2 values were 0.98 and 0.99.
119
A B LacZ activity (Modified Miller units) 100 140 WT C C+ WT R C+ 80 120 Cla35 R C+ Cla35 C C+ 100 60 80
60 40 C+ background 40
20 20 LacZ activity(Modified miller units) 0 0 20 40 60 80 100 120 140 0 0 0.2 0.4 0.6 0.8 1 1.2 C - background OD 600 nm
120 Figure 7. (A) Beta-galactosidase assay Esp19 mutants in C+ (y-axis) and C- (x- axis) backgrounds; Esp19-pvuIIC (open triangles), Esp19-pvuIIR (closed triangles) and WT pvuIIC (open circles), and pvuIIR (closed circles) represent in- frame lacZ fusions. (B) Esp33-pvuIIC (open triangles), Esp 33 -pvuIIR (closed triangles) and WT pvuIIC (open circles), and pvuIIR (closed circles) represent in- frame lacZ fusions. Specific activities were determined via linear regression plotting modified Miller units versus culture density with R2 values of 0.99 and
0.98 respectively and values were compared to WT.
121
A B
LacZ activity (Modified Miller Units) LacZ activity (Modified Miller units) 250 140
120 200
100 150 80 100
C+ background 60 C+ background 40 50
20 0 0 20 40 60 80 100 120 140 0 50 100 150 200 250 C - background C - background
122 Figure 8. Both A and B are correlograms, with pvuIICR transcribed from plac.
The X-axis represents the pvuIIC (WT and mutant) lacZ fusions and Y-axis reprents the pvuIIR (WT and mutant) lacZ fusions. Empty symbols represent no pvuIIC in-trans and filled symbols represent C.PvuII provided in trans from other
compatible plasmid. The measure of specific activity along with standard error is
shown along the dotted line (slope) for the WT and the mutants.
123
(A) (B) 140 200 WT R WT R Cla35 R 120 Cla35 R Esp 19 R Esp 19 R Esp33 R 150 WT R C+ Esp33 R 100 Cla35 R C+ Esp19 R C+ Esp33 R C+ 80 (modified Miller units)
(modified Miller units) 100
60
40 50
20 R frame LacZ activity R frame LacZ activity 0 0 0 10203040506070 0 102030405060 C frame LacZ activity (modified Miller units) C frame LacZ activity (modified Miller units)
124 Figure 9. Relative mRNA levels for strains carrying WT pvuIIR and its variants. mRNA levels were measured using the SYBR green method. Quantitation was performed using a stardard curve after normilazation to recA. Plotted values were derived by normalizing each variant to the WT.
125
2
1.5
1
0.5
0 WT Esp19 Cla35 Esp33
126 Figure 10. (A) & (B) represent alternative hairpins with Arm1 ( ) and Arm 2
( ) preceding the RBS and ATG of pvuIIR. (B) In- frame deletions of both
Arm1 and Arm 2 were created and lacZ fusions were made to the pvuIIR frame.
Beta-galactosidase assays were performed and compared to WT pvuIIR (closed circles) fusions.
127
A AC U U C A A A A G A A A A U C U C U U GC U U G A G A U A A A C A A G A U C U A G U UG G U U U G AC U U U AA C U U A A A G A C G U C A U A U -6.0 kcal/mole A U A A -4.5 kcal/mole A C A G 1087 1042 1055 1021
B
ATCGATAGACAACATTGAGCGAATAGCAAATGCCTTAAATGTTTCTATA ClaI pvuIIR RBS TCAATACTAATGATGGAACACGAAAATGAGTCACCCAGATCTAAATAAA pvuIIR start BglII pvuIIC stop
C
y = -8.0084 + 94.475x R= 0.99666 WTRTFC+ WTRTFC- y = -8.275 + 85.489x R= 0.98377 ARMIDELC+ y = -0.49998 + 58.531x R= 0.99148 ARMIDELC- y = -4.3143 + 85.619x R= 0.98243 ARMIIDELC+ ARMIIDELC- y = -3.2288 + 59.143x R= 0.98869 y = -1.1867 + 60.397x R= 0.99431 90 90 80 80 70 70 60 60 activity 50 activity 50
LacZ 40 LacZ 40
30 30
20 20 10 10 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 OD 600 OD 600 nm nm
128 Table1. Summary of Beta-galactosidase assays
Variants C.PvuII lacZ reading frame R/C fused to in trans* pvuIIC pvuIIR Ratio lacZ Sp Activity nb SE a Sp Activity n b SEa
WT C+ 61 3 13 57 3 2.5 .93
WT C- 50 3 7 57 3 5 1.14
Cla35 C+ 14 2 3 17 2 0.5 1.21
Cla35 C- 10 2 --- 12 2 --- 1.2
Esp19 C+ 50 2 1.37 40 2 1.25 .8
Esp19 C- 34 3 1 43 2 --- 1.26
Esp33 C+ 41 2 1.75 194 2 9 4.7
Esp33 C- 43 3 1.2 122 3 4.9 2.8
129 Table2. Hairpin deletion
Mutation C.PvuII in trans* lacZ reading frame pvuIIR
Sp. Activity N b SEa
WT Arm 1 ∆ C+ 74 2 ---
WT Arm 1 ∆ C- 51 3 ---
WT Arm 2 ∆ C+ 68 2 --- C- 53 2 --- WT Arm 2 ∆
( *) C.PvuII protein provided in trans on another compatible plasmid.
(a) Indicates standard error
(b) Indicates the number of times the experiment was performed
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142 DISCUSSION
Expression of genes in E coli is subject to both transcriptional and translational controls. Interactions between proteins and RNA, RNA and RNA or
DNA and proteins regulate the gene expression either positively or negatively.
E coli Hfq protein was defined for the first time for its involvement in Qβ- phage
replication (Fernandez, et al., 1968; Fernandez, et al., 1972). During stress E coli
Hfq mutants show reduced expression of certain genes because Hfq-mediated
RNA- RNA interactions are affected (Storz et al., 2002; Repoila, et al., 2003).
Several studies on Hfq describe this doughnut-shaped Sm-like protein as the
central player for post-transcriptional gene regulation.
Hfq facilitates base-pairing between noncoding (nc) RNAs and their
targets, but how it does so is not clearly understood. Additional work by Lease
and Woodson (2004) and personal communication (Miculecky PJ and Feig A L) indicates that Hfq, DsrA and rpoS mRNA form a ternary complex. Such complexes were also implicated with co-immunoprecipitation experiments
(Zhang, et al., 2002). This raises the possibility that more than one RNA binding site on Hfq is required to promote base-pairing interactions between DsrA and rpoS mRNA.
We used site-directed mutagenesis to probe the interaction between Hfq, ncRNA and mRNA in vivo. Our collaborators investigated the in vitro interaction.
143 Mutations were selected on the basis of structural and phylogenetic information.
The mutants were grouped into three categories, central cavity mutations (Y55A,
K56A and H57A), those that involve the proximal face (Q8A, D9A, D40A, Q41A,
F42A and I59A) or those on the distal face (Y25X, 130X, and Q53A). Mutations
at these sites showed different effects on in vivo and in vitro. Previous work had shown reduced stability of DsrA RNA when Hfq is absent (Sledjeski, et al., 2001).
This effect was suggested to be due to the overlap of the Hfq binding and RNase
E cleavage sites on DsrA RNA.
The DsrA RNA half-life experiments demonstrated that Hfq Y55A behaves like wild type Hfq whereas Hfq Y56A did not provide any protection to DsrA from
the RNase E effects and degraded very rapidly. DsrA degraded in two phases,
90% of this degraded in the fast phase and the remaining 10% of the RNA
stayed a little longer. Although Y55A stabilized DsrA in vivo, it was not able to
interact correctly with rpoS mRNA. We did not see any noticeable effect either in
vivo or in vitro effects with proximal face mutations, some of which failed to accumulate in vivo also did not complement the hfq− strain. However the distal
face mutations like Y25D and I30D complemented the rpoS activation to wild
type Hfq in vivo and showed effective binding to DsrA and rpoS mRNA 5’ UTR in
vitro.
In summary, our in vivo findings along with in vitro results revealed that
Hfq possesses at least two separate binding surfaces. In additions findings form
144 our collaborators showed that distal face mutations affected the A27 binding. Poly-
(A) RNA binds the back face of the torus which could separate both structurally and spatially the effects of Hfq on polyadenylation from its base-pairing effects.
Continuing to explore post-transcriptional regulatory mechanisms investigated the PvuII type II RM system. In bacteria such as E coli, genes are usually expressed from polycistronic mRNA. Ribosomes from an upstream gene can reinitiate translation at the next frame by translational coupling. Reinitiation efficiency can vary, but is seen to increase when the distance between the termination and restart codon decreases (Schoner, et al., 1986).
PvuII type II RM system is derived from Gram negative bacterium Proteus vulgaris (Gingeras, et al., 1981). Work from our laboratory by Tao et al., (1991),
Toa et al., (1992), Vijesurier et al., (2000) revealed the role of C.PvuII in transcriptional regulation of its own gene and downstream endonuclease gene.
Studies from other labs demonstrated regulatory roles for orthologous C proteins in other type II RM systems including BamHI, BglII, AhdI, SmaI, Eco72I, Esp
1396I, BcsI and more. The C gene precedes the R gene, and the initiator of the
REase gene and the termination codon of the C gene are generally close and sometimes even overlap. In Esp1396I, (Cesnaviciene et al 2003) in addition to transcriptional regulation C-gene affects the translation of its own gene as well as the down stream REase gene via translational coupling.
145 Results from these studies encouraged us to investigate whether similar regulatory mechanisms existed in the PvuII RM system. So I prepared lacZ translational fusions both in the R frame and in the C frame. To distinguish transcriptional from translational effects, these fusions used a C.PvuII- independent promoter (Plac) in place of the native promoter. I also analyzed the two in-frame mutants (Esp19 and Esp33) and one frameshift mutant (Cla35) by measuring their beta-galactosidase activity.
The expression levels from assays on WT and in-frame insertion mutations showed no difference with lacZ fusions to the pvuIIR frame or pvuIIC frame or with or without C.PvuII being provided in trans. However, a frame shift mutation dropped levels of expression considerably in both frames, though again providing C.PvuII in trans had no effect. These results are consistent with translational coupling. However, we have not proven if R is still transcribed in a nonsense mutation and polarity is a possible explanation. In addition, moving the pvuIIC termination codon relative to the pvuIIR initiator had no apparent effect which also indicates that this effect was not an artifact of the shifted position of stop codon.
I also examined predicted alternative hairpins upstream of pvuIIR, one of this would occlude the translation initiation region. However deletion of hairpin arms had no apparent effect.
146 Conclusions
The in vivo and in vitro findings reveal that Hfq contains at least two distinct RNA- binding surfaces.
Based on my results, pvuIIR expression is translationally regulated by translation of pvuIIC. This translation control involves coupling.
147 Future work
1. Analyzing the proximal face mutations of Hfq for their affect on A27
binding.
2. Characterize rpoS mRNA 5’ binding sites on the torus of Hfq.
3. Measuring the transcript levels of the PvuII fusions, to determine whether
the pvuIIR gene is being transcribed or not in these mutants, where
regulation by translational coupling seems to occur.
4. Characterizing the extent of translation control of pvuIIC gene on pvuIIR
gene by making nested deletions that move the relative positions of the
pvuIIC terminator and pvuIIR initiator.
5. Including the whole ORF of REase and measure expression and follow
the pattern and change in the REase activity.
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185 ABSTRACT
In bacteria which lack a nucleus, transcription and translation are concurrent and the majority of gene regulation occurs at the transcriptional level.
However, regulating translation can yield very rapid responses, and there is increasing evidence of its role in bacteria.
Hfq, an RNA-binding chaperone protein is a global regulator that binds the regulatory RNA DsrA and facilitates its interaction with rpoS mRNA.
This interaction controls translation of rpoS, which specifies an important transcription factor. The present study explored the possibility that Hfq needs more than one binding site to make such interaction possible. Through mutational analysis of hfq, the in vivo interactions between Hfq, DsrA and rpoS mRNA were studied.
The role of cis-acting translation control signals was also explored. The
PvuII type II RM was studied to understand the role of translation control in a system requiring tight, time-dependent regulation. It was found that pvuIIC translation regulates the translation of pvuIIR and this control involves coupling.
186