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12-2010 THE GENETICS OF METAL UPTAKE IN CAULOBACTER CRESCENTUS: CHARACTERIZATION OF REGULATORS AND IDENTIFICATION OF RECEPTORS Leah Howell Clemson University, [email protected]
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THE GENETICS OF METAL UPTAKE IN CAULOBACTER CRESCENTUS: CHARACTERIZATION OF REGULATORS AND IDENTIFICATION OF RECEPTORS
A Dissertation Presented to the Graduate School of Clemson University
In Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy Genetics
by Leah Williamson Howell December 2010
Accepted by: Dr. Harry Kurtz Jr., Committee Chair Dr. Julia Frugoli Dr. Amy Lawton-Rauh Dr. Kerry Smith
ABSTRACT
Caulobacter crescentus is an oligotrophic, Gram negative bacterium with a
dimorphic lifecycle. It is a model system for cell cycle regulation and cellular
development. Since C. crescentus resides primarily in freshwater, this
microorganism can also serve as a model system for nutrient uptake by
oligotrophic freshwater species. This study focused on TonB-dependent
receptors and metal-dependent regulators in C. crescentus.
To maintain appropriate levels of iron within the cell, bacteria have
evolved mechanisms to control iron uptake, storage, and use. In microbes, there
are two protein families possessing members that accomplish this: Fur and Rrf2.
In the C. crescentus genome, there are two putative members from each family. I
conducted a phylogenetic analysis to better understand the role of these four
proteins. The Fur family proteins were most closely related to regulatory proteins
that sense manganese (Mur-like) and zinc (Zur-like). In the Rrf2 family, one
protein shared homology with IscR, which regulates Fe-S synthesis and utilization pathways, while the other protein apparently represents a new member of Rrf2 family of regulatory proteins. I found that this protein is produced in
response to metal limitation. Future work on this complex topic will focus on
determining which metal or metals each of these regulatory proteins responds to,
as well as determining the binding sites.
ii TonB-dependent receptors are responsible for transporting ferric- siderophore complexes into the periplasm of the cell. I was able to reduce the potential number of TonB-dependent receptors that might possess iron regulation to nine. However, results from another laboratory likely expand that number to thirteen. I found that C. crescentus is capable of utilizing at least five siderophores. Deletion analysis of three receptors, CC0028, CC1666, and
CC3336, did not allow for linking a specific complex to a receptor. However, I found that CC0028 and CC3336 transcripts were up regulated 2.5 and 2.0 fold respectively in iron limited conditions. Interestingly, CC3336 is apparently regulated by both iron and carbon starvation. With one possible exception, this is the only TonB-dependent receptor seen to be regulated by two different stressors, one of which has been shown to utilize a sRNA.
iii DEDICATION
My loving husband, Jonathan Howell, has been by my side in good times and bad throughout my graduate school career. I could never have done this without him, and I want to dedicate this dissertation to him. I also want to dedicate this to my parents, Charlie and Gina Williamson, who have always loved me, encouraged me, and believed in me. Without them, I would never have been confident enough to take my education this far.
iv ACKNOWLEDGEMENTS
I would like to thank my advisor, Dr. Harry Kurtz, who has been a truly great mentor. He has always believed in me and practiced extreme patience. He was always there to answer questions, and his guidance throughout this project was crucial.
I would like to thank my committee members, Dr. Julia Frugoli, Dr. Kerry
Smith, and Dr. Amy Lawton-Rauh. Their suggestions and guidance were very helpful to me and always insightful.
I would like to acknowledge Amy Lawton-Rauh and Brad Rauh for their valuable assistance on the phylogeny work in this dissertation.
I would also like to thank the present and past members of the Kurtz lab who were often very helpful and supportive.
Finally, I would like to express gratitude to my family and friends for all of their love and support during my graduate school career. I am truly lucky to have such wonderful people in my life.
v TABLE OF CONTENTS
Page
TITLE PAGE ...... i
ABSTRACT...... ii
DEDICATION...... iv
ACKNOWLEDGMENTS ...... v
LIST OF TABLES...... viii
LIST OF FIGURES ...... ix
CHAPTER
I. LITERATURE REVIEW ...... 1
Introduction...... 1 The biological significance of iron...... 2 Iron toxicity ...... 5 Other important metals...... 7 Metal-dependent regulation...... 8 Iron uptake ...... 15 Environmental iron levels ...... 19 Purpose of research ...... 23
II. METAL-DEPENDENT REGULATORS IN C. CRESCENTUS ...... 35
Introduction...... 35 Methods...... 40 Results ...... 45 Discussion ...... 56
III. TONB-DEPENDENT IRON UPTAKE IN C. CRESCENTUS...... 65
Introduction...... 65 Methods...... 69 Results ...... 90 Discussion ...... 106
vi Table of Contents (Continued)
Page
IV.SUMMARY AND FUTURE DIRECTIONS ...... 114
Introduction...... 114 Metal-dependent regulators...... 115 Iron-regulated TonB-dependent receptors...... 120 Conclusion...... 125
vii LIST OF TABLES
Table Page
1.1 Several proteins that require iron to function properly ...... 4
2.1 Fur family pairwise comparison ...... 46
2.2 Rrf2 family pairwise comparison ...... 50
3.1 Primers used to construct and test for deletions ...... 72
3.2 Primer pairs used for RT-PCR and for RNA slot blots for each ORF ...... 78
3.3 Slot blots to confirm the iron regulation of TonB-dependent receptors ...... 100
3.4 Summary of findings from three experiments ...... 101
viii LIST OF FIGURES
Figure Page
1.1 The action of Fur at the Fur box of target genes...... 10
1.2 Model of TonB-dependent transport of iron ...... 17
1.3 Representation of the amount of iron available in the North Atlantic and North Pacific in colloidal iron and free iron forms ...... 20
1.4 Concentration of free iron in metal polluted, eutrophic, and mesotrophic freshwater sources...... 22
2.1 The action of Fur at the Fur box of target genes...... 36
2.2 Bayesian Fur family phylogeny ...... 47
2.3 Maximum Likelihood Fur family phylogeny ...... 48
2.4 Bayesian Rrf2 family phylogeny...... 51
2.5 Maximum Likelihood Rrf2 family phylogeny...... 52
2.6 Outer membrane protein profile of CB2A cells grown in different media...... 54
2.7 Western blot of HIS-tagged CC2625 ...... 55
3.1 Number of annotated TonB-dependent receptors in the genomes of several microorganisms...... 67
3.2 SacB counter-selection...... 71
3.3 CC0028 gene deletion strategy ...... 73
3.4 CC1666 gene deletion strategy ...... 74
3.5 CC3336 gene deletion strategy ...... 75
3.6 Outer membrane profile and MS/MS data for CB2A...... 91
ix List of Figures (Continued)
Figure Page
3.7 Growth at OD600 of two C. crescentus strains on FeCl3 ...... 92
3.8 Growth at OD600 of two C. crescentus strains on ferric-rhodotorulate ...... 93
3.9 Growth at OD600 of two C. crescentus strains on ferric-citrate ...... 94
3.10 Growth at OD600 of two C. crescentus strains on ferric-DHB ...... 95
3.11 Growth at OD600 of two C. crescentus strains on ferric-ferrichrome ...... 96
3.12 Growth at OD600 of two C. crescentus strains on ferric-enterobactin...... 97
3.13 Reverse transcriptase PCR (RT-PCR) to identify iron regulated TonB-dependent receptors...... 99
3.14 Deletion confirmations through PCR...... 102
3.15 Map of 1666 deletion in CB15...... 103
3.16 Growth of CB15 CC3336 deletion mutant on various concentrations of iron and carbon ...... 105
4.1 The [2Fe-2S] coordinating site of IscR as comparison for other proteins ...... 118
4.2 Two possible scenarios for the regulation of CC3336...... 122
x CHAPTER ONE
LITERATURE REVIEW
Introduction
Caulobacter crescentus is a gram negative α-Proteobacterium that inhabits nutrient poor freshwater (1, 22, 37, 40, 56, 90). Because this microbe has a dimorphic life cycle, it is often used as a developmental model (42, 43, 50,
56, 78, 95). The prosthecate, or stalked cell, is the sessile form of C. crescentus development that uses an adhesive holdfast at the distal tip of the stalk for surface attachment (54, 56, 75, 78, 87). The stalked cell grows a flagellum at the pole opposite to the stalk and then divides, creating a swarmer cell and leaving the stalked cell to divide again (5, 21, 42-44, 54, 73). The swarmer cell is motile and actively seeks out nutrients but does not have the ability to replicate its DNA
(21, 42, 95). Ultimately, the swarmer cell will become a stalked cell when it loses its flagellum and attains a stalk, often attaching to a surface and gaining the ability to replicate its chromosome (21, 43, 73, 75, 78). The stalk is an extension of the cell that contains no DNA, ribosomes, or nucleoplasm (40, 75). It has an adhesive holdfast at the end to adhere to surfaces, creating a very strong attachment (22, 78). C. crescentus also has the notable ability to live in extremely nutrient dilute conditions, and can even be found in commercially sold bottled water (1, 37, 38, 64). Because many studies have examined this microorganism as a model for life cycle and cell cycle regulation, there are many molecular tools
1 available for the study of C. crescentus. There is currently no model for metal
uptake in freshwater microbes. With the variety of tools available and the ability
to live in extremely oligotrophic environments, C. crescentus is a good candidate model for nutrient uptake in freshwater microorganisms.
The biological significance of iron
Billions of years ago in Earth’s anaerobic atmosphere, microbes developed mechanisms for iron homeostasis that functioned under the environmental conditions of the time (2, 47, 100). The main form of iron available was ferrous iron (Fe2+) which is more soluble than ferric iron (Fe3+) (2). Iron used
to be a biologically “friendly” metal, and organisms evolved to use it for many
cellular processes when oxygen was not present at high concentrations (2). With
the beginnings of oxygenic photosynthesis, O2 levels rose in the environment,
resulting in the conversion of ferrous iron to ferric iron, a form of iron less readily
available to microorganisms (2). Iron is one of the most abundant elements in the
Earth’s crust which is why it made sense for early organisms to adapt its use to
so many cellular functions (2, 53). Oxygen is now present at high levels on Earth
and is present in the environments of many microbes, making it difficult for
organisms to acquire enough iron due to the insolubility of ferric iron minerals (2,
100). Additionally, in an aerobic system, too much cellular iron can be toxic to
organisms through the formation of free radicals (2, 47). Therefore, microbes
2 must carefully balance the amount of iron in the cell, as too much is just as
detrimental as too little.
Under aerobic conditions, the primary form of iron available, ferric iron, is
present in colloidal ferric oxyhydroxides and compounded with dissolved organic molecules, except in low pH environments where it is much more soluble as a free ion (32, 47, 53, 59, 86). Free ferric iron has a solubility of just 10-18 M at
physiological pH, making it difficult for organisms to obtain enough (2, 24, 28, 47,
53, 86). Iron is needed in many different pathways including oxygen activation,
biosynthesis of amino acids and nucleosides, and gene regulation (2, 10, 31, 47,
57, 69, 70, 84, 86, 107). Most importantly, it plays a major role in cellular
respiration and photosynthesis as a component of electron transport systems
where it is incorporated into cytochromes and iron-sulfur proteins (2, 57). Iron’s
use in the cell depends almost exclusively on its integration into proteins (2). It is
a useful cofactor for many enzymes due to its ability to lose and gain electrons
with relative ease, switching from Fe(II) to Fe(III) (12, 47, 69). Several proteins
that require iron in order to properly perform their cellular functions are listed in
Table 1.1. Although having enough iron is crucial, having more than enough iron
is toxic due to the formation of oxygen and hydroxyl radicals (23, 47, 59, 69).
Therefore, the employment of a system for regulating intracellular iron levels is critical. Iron regulation occurs at the levels of uptake, storage, use, and through the detoxification of harmful radicals (2, 3, 47).
3 Table 1.1: Several proteins that require iron to function properly. Included are examples of proteins from various important cellular processes.
General process Specific process Protein Catabolism electron transport chain succinate-coenzyme Q reductase (67)
cytochrome bc1 complex (14) Terpredoxin (60) Krebs cycle Aconitase (51) electron transport chain and Krebs cycle Succinate dehydrogenase (67) methanol catabolism formate dehydrogenase (76)
Anabolism amino acids glutamate synthase (98) Nitrogenase (82) Pyrimidine-deoxynucleoside 1'-dioxygenase nucleotides (96) ribonucleotide reductase (11)
Oxygen radical superoxide neutralization neutralization superoxide dismutase (88) hydrogen peroxide neutralization catalase (88) peroxidase (88)
iron-dependent Gene regulation regulation Fur (16) RirA (12) IscR (30)
4 Iron toxicity
Hydrogen peroxide and superoxide toxicity are common problems, and bacteria grown in excess iron are often found to have increased sensitivity to these redox stress agents (2). Hydrogen peroxide is formed when superoxide interacts with itself, and superoxide is a reduced form of oxygen (2, 59, 111).
Superoxide and hydrogen peroxide are not as harmful as hydroxyl radicals, but they still cause damage to cellular components (2, 57). The presence of excess iron in the cell can lead to interaction with reactive oxygen species through such processes as iron reduction (Equation 1) and the Fenton reactions (Equation 2)
(2, 12, 47, 69, 86, 102, 111). These pathways add up to the Haber-Weiss reaction (Equation 3) and lead to the production of hydroxyl radicals, an extremely harmful byproduct (2, 47, 59, 69, 86). Hydroxyl radicals interact with and break down lipids, proteins, and DNA (35, 57, 59, 99, 111).
-3+2+ Equation 1: O2 +Fe Fe +O2
2+ 3+ - Equation 2: Fe + H2O2 Fe + OH + HO.
-- Equation 3: O2 +H2O2 HO. + OH + O2
Enzymes involved in free radical neutralization have evolved to protect
cells from damage; these include superoxide dismutases (SODs), catalases, and
5 peroxidases (2, 47, 57, 59, 88, 92). These enzymes neutralize superoxide and
hydrogen peroxide, and they have been found in many bacteria (57, 59, 92).
SOD has been found in every aerobic microbe studied, and its low abundance in
obligate anaerobes is thought to be the basis for oxygen’s toxic effect on these organisms (57). However, there are also other reasons for the toxic effect of oxygen including the inhibition of nitrogenase and other enzymes (82). Catalase and peroxidase both neutralize hydrogen peroxide (Equations 4 and 5), although peroxidase requires the addition of a reductant (often NADH) (57). Both enzyme types (SODs and catalases/peroxidases) require metals to work properly. SOD, which catalyzes the degradation of superoxide (Equation 6), uses divalent metals in order to function; these include manganese, iron, and copper-zinc (48, 92).
The combined reaction of catalase and SOD to neutralize radicals can be seen in
Equation 7. Caulobacter crescentus contains both an iron (FeSOD) and a copper-zinc SOD (CuZnSOD) (41, 88, 91-93). The copper-zinc SOD is active in the periplasm and protects against superoxide outside the cell while the iron form of SOD acts in the cytosol (41, 88, 92). C. crescentus has also been shown to have catalase activity, and the bifunctional catalase-peroxidase gene has been identified, although little is known about the enzyme in this organism (41, 88, 94).
Catalases and peroxidases, when studied in various other organisms, have required metals to work, generally in the form of heme or manganese (6, 48,
111).
6 Catalase Equation 4: H2O2 +H2O2 2H2O+O2
++Peroxidase Equation 5: H2O2 +NADH+H 2H2O+NAD
Superoxide dismutase -- + Equation 6: O2. +O2. +2H H2O2 +O2
Superoxide dismutase & -+. Catalase Equation 7: 4O2 +4H 2H2O+3O2
Another way that bacteria deal with having excess iron in the cell is by storing it for times when it is needed. Bacterioferritins are intracellular proteins used to store excess iron for times of iron starvation (2, 3). One major difference between ferritins and bacterioferritins is that bacterioferritins possess a heme group while ferritins do not; both molecules can be found in many bacteria, though bacterioferritin is found more often than ferritin (2, 3). Bacterioferritin proteins form a spherical shell in which up to 4000 iron atoms can reside, and the shell helps to avoid loss of iron until it is released in times of iron starvation (3).
Other important metals
There are other metals besides iron that are micronutrients required for bacterial growth. Manganese (Mn) is used as a cofactor for certain radical detoxifying enzymes such as catalases and peroxidases and is a cofactor for
7 various other enzymes, including those involved in metabolic processes such as water oxidation in photosynthetic bacteria (48, 57). Manganese can change oxidation states, so it is a good cofactor for reducing enzymes (48). However, it cannot switch between oxidation states as readily as iron and, therefore, has less reducing potential (48). Molybdenum (Mo), another essential metal to most living systems, is present in various enzymes where it is coordinated with iron or tungsten and employed as a cofactor for reactions in the metabolism of carbon, nitrogen, and sulfur (36, 49). Zinc (Zn) is crucial to all living things and is the only metal that has a representative enzyme in every essential enzyme class that is currently known (71). It is highly important in many cellular processes including being used as a structural component of transcription factors (57, 71). Cobalt,
(Co) is also biologically important, because it is a component of cobalamin (B12) and used for the activity of various enzymes including methylmalonyl-CoA mutase which uses cobalamin as a cofactor for carbon rearrangements (29, 57).
Overall, maintaining a good concentration of metals in the cell is crucial to survival.
Metal-dependent regulation
Due to the issue of iron’s insolubility and toxicity in aerobic environments, bacteria have developed systems to tightly regulate iron uptake, storage, and use. The ferric uptake regulator (Fur) protein mediates iron-dependent expression in many bacteria including Escherichia coli and is considered the
8 global regulator of iron-dependent genes (19, 23, 45, 86). Fur has been shown to
repress over 100 genes in E. coli and Pseudomonas aeruginosa (2, 19, 52, 102).
This protein functions as a dimer consisting of two ~17 kD monomers (24, 86).
Fur has an N-terminal DNA binding region and a C-terminal dimerization domain which makes Fur a dimer regardless of whether or not iron is bound (17, 24, 52,
86). Each monomer has an iron and a zinc ion binding site, and binding of Fe2+ causes a conformational change in the N-terminal DNA binding region, allowing
Fur to interact with specific sequences of DNA called Fur boxes (17, 52). The Fur box is an ~19 bp AT-rich palindromic sequence located in the promoter of Fur- regulated genes (17, 26, 27, 34, 35, 52). Fur binding at the Fur box represses target genes (8, 24, 52, 86). When iron is depleted or present at only low amounts, Fur does not bind iron as well, and Fur-regulated genes are derepressed (24, 27, 52, 83). Iron loaded-Fur binding at the promoter, typically between the -10 and -35 sites, hinders RNA polymerase binding, thus halting transcription as shown in Figure 1.1 (2, 17, 52, 70). Fur also has the ability to induce expression of genes in response to adequate iron in E. coli and
Helicobacter pylori (2, 17, 18). These are genes that require up-regulation in
times of adequate iron, such as iron-containing enzymes and iron-storage
proteins. An indirect mechanism is used where Fur represses small regulatory
RNAs (sRNAs) that would normally repress these genes, so Fur is still acting as
a repressor in this pathway (2, 18, 52, 66, 102).
9
Figure 1.1: The action of Fur at the Fur box of target genes. Arrow indicates transcription start site (+1). A: the Fur dimer (purple) is always bound by zinc, a structural ion.
When iron levels are insufficient, Fur cannot bind with Fe2+ and loses affinity for the Fur box. This allows RNA polymerase to bind and initiate transcription. B: iron is at sufficient levels to bind Fur.
This changes the confirmation of Fur so it binds to the Fur box. This blocks RNA polymerase from initiating transcription.
10 The manganese uptake regulator (Mur) was first discovered in R. leguminosarum (4, 18, 19, 45, 101). Mur shares sequence homology with Fur and can even bind to E. coli Fur boxes and complement Fur mutants (19, 45, 84).
In response to sufficient manganese levels, Mur negatively regulates the sitABCD operon, which encodes an ABC-type transporter for Mn2+ uptake (4, 18,
19, 45, 84, 101). The promoter binding site for Mur is called the MRS and has a different sequence than the Fur box, but it is similar to the Fur box in that it is palindromic and ~19 bp (4, 19, 45, 84). Manganese and iron dependent regulation are often interchangeable, and high concentrations of manganese interfere with iron homeostasis (79). This illustrates the reason why manganese regulation is so important. In organisms other than rhizobia, the manganese uptake regulator is MntR, which is unrelated to the Fur family and is a DxtR family member (45, 74).
The iron response regulator (Irr), also a member of the Fur family, has been characterized in Bradyrhizobium japonicum (4, 19, 84, 101). Irr is present in all Rhizobials and Rhodobacterales but has not been found outside this group of organisms (18, 45). Irr acts by repressing or activating gene expression in iron insufficient conditions by binding to ICE sequences near the promoter; it negatively regulates iron-containing proteins and positively regulates genes for high-affinity iron uptake (24, 45, 79, 84, 101). Irr is degraded when it binds heme, so in iron replete conditions, Irr levels decrease in the cell (4, 24, 45, 79, 84,
101). In iron deficient conditions, Irr represses genes for heme biosynthesis
11 among others (4, 24, 84, 101). Recently, Irr in B. japonicum was shown to bind manganese when intracellular iron is limiting and prevent heme from binding
(79). This stabilizes Irr under iron limited conditions (79).
The zinc uptake regulator (Zur) was found first in E. coli and Bacillus subtilis (4, 15, 24, 39). It is another Fur family member which acts by repressing genes for zinc uptake (4, 24, 39, 97). It does so in response to high Zn2+ levels and represses genes for the zinc ABC transport system, helping to maintain intracellular zinc homeostasis (4, 39). The Zur DNA binding site is very similar to the Fur box and is called the Zur box (24, 39). Unlike iron, in physiological conditions, zinc does not go through reduction or oxidation (85, 105). However, zinc is still crucial to the activity of various enzymes and is incorporated into proteins required for cell maintenance and survival (85, 105). An overabundance of zinc can cause this metal to take over the active site in non-zinc proteins and lead to the generation of reactive oxygen species (85, 105). Like iron, having excess cellular zinc levels creates potentially toxic conditions; this explains why there is tight regulation on zinc uptake like there is with iron (15, 24, 39, 97).
In addition to the Fur family of proteins which includes Fur, Mur, Irr, and
Zur, another family of proteins with known iron-dependent regulators is the Rrf2 family. The rhizobial iron regulator (RirA), the iron-sulfur cluster regulator (IscR), and NsrR are all members of the Rrf2 family of proteins (2, 45, 101, 103). These three proteins are the only well studied members of this protein family (45, 101).
RirA from α-Proteobacteria such as R. leguminosarum and S. melitoti is a protein
12 that has also been shown to regulate iron-responsive genes and performs a
similar job to the Fur protein (2, 4, 18, 84, 102, 104, 110). RirA was first identified in Rhizobium, hence the name RirA (rhizobial iron regulator) (45, 101). In organisms such as Rhizobium and Sinorhizobium, RirA has been shown to bind to iron-responsive operators (IROs) at the promoters of specific iron-regulated genes; IROs have a very different characteristic sequence than Fur box sequences (45, 84, 110). The IRO is not palindromic, is typically shorter than the average Fur box at ~17 bp long, and it does not share sequence similarity with the Fur box (84, 110). RirA binds to IROs in the promoter and represses transcription when sufficient iron is available (110). This protein represses over
80 transcriptional units in response to iron in Rhizobium and Sinorhizobium (12,
45, 101). It is believed that because it can bind [2Fe-2S] clusters like its Rrf2 relative, IscR, RirA is binding DNA in response to an Fe-S cofactor in order to regulate gene expression (45).
IscR is also a member of the Rrf2 family and is a Fe-S cluster biogenesis gene regulator in E. coli (2, 30, 83, 89, 101, 110). Fe-S cluster containing proteins are widely seen in biology (30, 89). These clusters are used for processes such as electron transfer, structural stability, and transcriptional regulation (69). IscR regulates genes when it binds [2Fe-2S] but has a relatively low affinity for these clusters compared to other proteins (2, 30). This is probably so that IscR only regulates genes when Fe-S clusters are available at a higher level than required for cellular homeostasis (2, 30). In response to high [2Fe-2S]
13 levels, IscR represses genes responsible for Fe-S cluster biogenesis (2, 30, 45,
101). When Fe-S clusters are at a sufficiently low level, genes under the control of IscR are transcribed (2, 30).
Another Rrf2 family member is NsrR, a nitrite-responsive regulator (12, 45,
103). This protein has been found in some Proteobacteria including E. coli where it regulates over 30 genes (45, 72, 103). NsrR is responsible for the regulation of genes that are involved in nitric oxide (NO) metabolism (12, 45, 81, 103). When
NO is at a high cellular level, genes that oxidize or reduce it need to be transcriptionally active (72, 81, 103). NO binds to NsrR, probably on an Fe-S cluster, to create a transcriptional response (72, 81, 103). This seems to confer a loss of DNA binding activity in NsrR, ensuring that genes involved in NO oxidation or reduction are transcribed, reducing the toxic effects of NO to the cell
(81, 103).
All of the regulators mentioned act similarly to Fur in reaction to adequate levels of their respective binding atom or molecule (Figure 1.1). The exceptions to this are Irr which can transcriptionally repress or activate genes and NsrR which loses the ability to bind DNA when NO is present (24, 45, 79, 84, 101).
Generally, all of these regulators are very much alike in action although binding sites for each are quite different.
14 Iron uptake
Little is known about iron uptake in freshwater microorganisms (16, 45).
Many gram negative bacteria employ a system that uses siderophores to tightly
bind ferric iron (Fe3+) and deliver it to high affinity TonB-dependent outer
membrane receptors (107). Siderophores are small molecules secreted by many
aerobes to help scavenge iron by acting as a chelator that solublizes ferric iron
(13, 25, 31, 32, 86, 108). Siderophores contain functional groups to bind ferric
iron such as hydroxamates, catechols, and α-hydroxycarboxylates (2, 53, 57, 84,
86). Some siderophores even employ multiple types of these functional groups
(53, 86).
Though ferric-siderophore complexes are soluble, they cannot simply
diffuse through the outer membrane, so they are transported into the cell through
high affinity TonB-dependent receptors (24, 33, 63, 86, 108). These receptors
are used to transport nutrients such as carbohydrates, vitamin B12 (cobalamin),
and chelated ferric iron (13, 32, 77). They are made of up a C-terminal β-barrel that transverses the outer membrane (OM) and a “cork” (2, 84). The “cork” is made up of a globular domain that takes up the first ~160 amino acids of the N- terminal region, and it “plugs” the periplasmic portion of the beta barrel (2, 84).
When a ferric-siderophore binds to the receptor, it causes a conformational change, allowing the siderophore complex to be imported into the periplasm (2,
32, 84). Each receptor imports only one type of siderophore and is highly specific, so bacteria generally have several receptors to be able to utilize several
15 different siderophores (2, 84). C. crescentus strain CB15 has 66 annotated
TonB-dependent receptors encoded in its genome (65). This is a high number
when compared to other bacteria that have been studied; only two of these 66
receptors have a known function, and both are involved in carbohydrate uptake
(20, 64, 65). One of the main ligands for TonB-dependent receptors is ferric iron
bound by siderophores, and these receptors are tightly regulated by iron
availability due to the harmful effects of having too much intracellular iron (2, 7).
In the inner membrane of many gram negative microbes, there is a complex of three proteins: ExbB, ExbD, and TonB (2, 77, 84, 86). As indicated in
Figure 1.2, this complex harnesses the electrochemical charge of the cellular
membrane (CM) in order to move the “cork” of a TonB-dependent receptor in the
OM, allowing ferric siderophores to enter the periplasm (2, 32, 84). ExbB and
ExbD are imbedded in the CM, while TonB is located in the periplasm with its N- terminal region imbedded in the CM (2, 77). TonB has a proline-rich extender
region and a C-terminal region that transmits energy to the N-terminal “TonB
box” of the TonB-dependent receptor in the OM (2, 77, 84, 86). It is thought that
ExbB and ExbD focus the energy of the CM into a TonB conformational change,
or an “energized” version of TonB, which has been shown to use its proline-rich
region to stretch and physically interact with the TonB box of a TonB-dependent
receptor, allowing the “cork” to open and import a ferric-siderophore into the
periplasm (2, 77, 84). In order to be transported across the CM, a periplasmic
binding protein (PBP) and ATP-binding cassette (ABC) transporter are employed
16
Figure 1.2: Model of TonB-dependent transport of iron. Extracellularly, ferric iron is bound by a siderophore. This complex is recognized by a receptor in the OM. TonB is in the
CM where ExbB and ExbD help it harness the energy of the CM and store it as a conformational change. The charged form of TonB interacts with the OM receptor. This allows the “cork” of the receptor to open, bringing the ferric-siderophore into the periplasm.
17 (2, 84, 86). The PBP shuttles the ferric-siderophore through the periplasm to an
ABC receptor in the CM which hydrolyzes ATP to pass the ferric-siderophore through the CM (2, 84, 86). Once inside the cell, the iron attached to the siderophore is thought to be reduced, releasing it from the siderophore, which can then be recycled (2, 84). Another way iron is released is through the breakdown of siderophores (84). Both siderophores and high affinity iron-uptake receptors are negatively regulated in response to increasing iron levels in all bacteria studied (53).
Though the stalk of the stalked C. crescentus cell is believed to be involved in nutrient uptake due to an elongation response under phosphate starvation, the ExbB protein cannot be found in the stalk region (106). This may indicate that the stalk is incapable of TonB-dependent transport as it is currently understood (106). However, the apparent absence of ExbB in the stalk may be due to experimental error such as the protein failing to focus well under isoelectric focusing (40). Overall, little is known about nutrient uptake in C. crescentus.
The types of siderophores that can be found vary greatly between different environments (58, 86). Marine siderophores characteristically consist of α- hydroxycarboxylate functional groups (9, 58, 86). They are typically amphiphilic and photoreactive when bound to ferric iron which is not generally the case for siderophores found in terrestrial environments or produced by pathogenic bacteria (9, 58, 86). Even though the typical structure and other characteristics of
18 marine siderophores is known, very few marine siderophores have been found
as compared to the number of terrestrial siderophores that are known (86).
Besides the composition of siderophores, little is known about iron acquisition in marine bacteria, and even less about freshwater organisms (58, 86).
Environmental iron levels
Terrestrial environments typically have much more available iron than aquatic habitats (58, 86). Aquatic environments also differ in their iron availability.
In oceans, more than 99% of dissolved iron is strongly bound by organic
compounds (32, 61, 109). One form of bound iron, colloidal iron species, most
likely decrease iron availability through the accumulation of colloidal iron into
larger particles which settle, decreasing readily available iron in the water column
(109). Particles with a molecular diameter of <0.02 µm are considered soluble
iron, while particles with diameter from 0.02-0.4 µm are considered colloidal iron
(109). A study by Wu et al. (109) looked at soluble and colloidal iron levels in the
North Atlantic and North Pacific oceans. Soluble iron, similar to phosphate and
nitrate levels, increases in relation to the depth of the water and ranges from ~0.1 nM at the surface to ~0.4 nM at a depth of ~1000 m (109). The opposite is true of colloidal iron, which has the highest levels at the surface of the ocean and was measured in levels ranging from ~0.02 to ~0.4 µM (Figure 1.3) (109).
19
Figure 1.3: Representation of the amount of iron available in the North
Atlantic and North Pacific in colloidal iron and free iron forms. Data in figure is from a study done by Wu et al. (109).
20 The concentration of dissolved iron in lake water is higher than the
concentration in seawater, and less iron is bound to organic compounds (61). In a study done by Nagai et al. (61) of Lake Kasumigaura in Japan and two rivers running into the lake, the two rivers had a mean dissolved iron concentration of
1.5 µM, while the iron concentration in the lake was 0.14 µM. This is most likely higher than typical freshwater because Kasumigaura is a eutrophic lake (61). A
1990 study of Lake Piaseczno, a mesotrophic lake in Poland, found a mean
dissolved iron concentration of 0.78 µM (80). In a 2008 study, Lake Beysehir, a
metal polluted lake in Turkey had a mean dissolved iron concentration of 14 µM
(62). Another study done in 2004 of an additional metal polluted lake in Turkey,
Lake Hazar, saw a mean dissolved iron concentration of 4.2 µM (68). As
presented in Figure 1.4, it appears from these few studies that dissolved iron
tends to reach average levels of up to about 1.5 µM in eutrophic freshwater
(average iron level in rivers of Kasumigaura) and about nine fold higher in a polluted body of water (Lake Beysehir). Freshwater, the environment in which C. crescentus dwells, is quite low in iron for the growth of many microbes, because bacteria tend to require about 1-100 µM iron to grow at an optimum level (2).
21 Lake Beysehir 14
Lake Hazar 4.2
Rivers of 1.4785 Kasumigaura
Lake Piaseczno 0.78
Lake 0.1445 Kasumigaura
0 2 4 6 8 10121416 Mean concentration of free iron (µM)
Figure 1.4: Concentration of free iron in metal polluted, eutrophic, and mesotrophic freshwater sources. Lakes Beysehir and Hazar are metal polluted (62).
Lake Kasumigaura and the associated lakes are eutrophic (61), and Lake Piaseczno is
mesotrophic (80).
22 Purpose of research
The goal of this study was to develop a model for iron-dependent uptake and overall regulation in C. crescentus. This freshwater microbe, as well as the rest of the order Caulobacterales, has a large number of annotated TonB- dependent receptors as compared with other organisms. C. crescentus also has been studied very little with respect to metal-dependent regulators. Most of the studies of high affinity iron uptake systems have centered on pathogens such as
E. coli, N. gonorrhoeae, V. cholerae and H. influenzae and soil microbes such as
B. japonicum and P. putida (2, 10, 25, 46, 53, 55, 77). Very little work has been done on freshwater bacteria in this area (32). C. crescentus is an interesting candidate for the study of iron regulation and iron uptake systems because there are many genetic tools available for the study of this microorganism, it has a large number of receptors, and it could possess a novel member of the Rrf2 family. The goal of this research was to answer the questions: (1) What are the key players in metal-dependent regulation in this microorganism? and (2) Which
TonB-dependent receptors are iron regulated?
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34 CHAPTER TWO
METAL-DEPENDENT REGULATORS IN C. CRESCENTUS
Introduction
Due to the issue of iron’s insolubility and toxicity in aerobic
environments, bacteria have developed systems to tightly regulate iron uptake,
storage, and use. The ferric uptake regulator (Fur) is responsible for iron-
dependent regulation in many organisms including Escherichia coli (13, 15, 25,
39). It is considered to be the global iron response regulator (13, 15, 25, 39). This
protein uses ferrous iron (Fe2+) as a cofactor in order to bind to the Fur box
where it mediates transcriptional regulation (6, 7, 11, 16-19, 28, 34, 35, 37, 39,
52). Iron bound-Fur binds at the promoter, typically between the -10 and -35 sites
and hinders RNA polymerase binding, thus preventing transcription (Figure 2.1)
(3, 11, 28, 32). Fur can also activate the expression of genes such as iron-
containing enzymes and iron-storage proteins in response to adequate iron in E.
coli and Helicobacter pylori (3, 11, 12). However, an indirect mechanism is used
wherein Fur represses the transcription of small regulatory RNAs that would
normally repress these genes, so Fur is still acting as a repressor in this pathway
(3, 12, 28, 31, 49).
The manganese uptake regulator (Mur) was first discovered in
Rhizobium leguminosarum (6, 12, 13, 25, 48). Mur shares sequence homology with Fur and can even bind to E. coli Fur boxes and complement Fur mutants
35
Figure 2.1: The action of Fur at the Fur box of target genes. Arrow indicates transcription start site (+1). A: the Fur dimer (purple) is always bound by zinc, a structural ion.
When iron levels are insufficient, Fur cannot bind Fe2+ and loses affinity for the Fur box. This allows RNA polymerase to bind and initiate transcription. B: iron is at sufficient levels to bind to
Fur. This changes the confirmation of Fur so it binds to the Fur box. This blocks RNA polymerase from initiating transcription.
36 (13, 25, 38). Mur regulates the sitABCD operon encoding an ABC-type transporter for Mn2+ uptake (6, 12, 13, 25, 38, 48). Mur represses transcription of this operon under Mn2+ sufficient conditions by recognizing a DNA promoter sequence called the MRS (6, 13, 25, 38).
The zinc uptake regulator (Zur) was found first in E. coli and Bacillus subtilis (6, 9, 18, 23). It is another Fur family member which acts mainly by repressing genes (6, 18, 23, 47). It does so in response to high Zn2+ levels and represses genes for the zinc ABC transport system, helping to maintain intracellular zinc homeostasis (6, 23). The Zur DNA binding site is called the Zur box (18, 23).
In addition to the Fur family of proteins, another family of metal-dependent regulator proteins is the Rrf2 family. The rhizobial iron regulator (RirA), the iron sulfur cluster regulator (IscR), and NsrR are all members of the Rrf2 family of proteins (3, 25, 48, 50). These three proteins are the only well studied members of this protein family (25, 48). RirA from α-Proteobacteria such as R. leguminosarum and Sinorhizobium melitoti is a protein that regulates specific iron-responsive genes and performs a job similar to the Fur protein (3, 6, 12, 38,
49, 51, 52). RirA was first identified in Rhizobium, hence the name RirA
(Rhizobial iron regulator) (25, 48). RirA binds to its target promoter DNA sequences, Iron-responsive operators (IROs), and represses transcription only when sufficient iron is available (25, 38, 52). A closely related Rrf2 family member, IscR, is an Fe-S cluster biogenesis gene regulator in E. coli (3, 20, 37,
37 41, 48, 52). In response to high [2Fe-2S] levels, IscR represses genes responsible for Fe-S cluster biogenesis, and when [2Fe-2S] is limiting, these genes are derepressed (3, 20, 25, 48). It is believed that because RirA can bind
[2Fe-2S] like its Rrf2 relative, IscR, RirA is probably also binding DNA in response to an Fe-S cofactor in order to regulate gene expression (25).
Another Rrf2 family member is NsrR, a nitrite-responsive regulator (8, 25,
50). This protein has been found in various Proteobacteria including E. coli (25,
33, 50). NsrR is responsible for the regulation of genes that are involved in nitric oxide (NO) metabolism (8, 25, 36, 50). NO binds to NsrR, probably on an Fe-S cluster, to create a transcriptional response (33, 36, 50). This confers a loss of
DNA binding activity in NsrR, ensuring that genes involved in NO oxidation or reduction are transcribed (36, 50).
All regulators mentioned act similarly to Fur in reaction to high enough levels of their respective binding atom or molecule. While these regulators are very much alike in action, the binding sites for each are quite different. With the exception of some relatively recent work on RirA in rhizobia, very little is known about iron dependent regulation in any of the α-Proteobacteria (25). Using an informatics approach, Rodionov et al. (37) showed that Caulobacter crescentus has two possible metal-dependent regulators including a putative Fur (though they looked for no other Fur family members) and a putative member of the Rrf2 family of proteins. Work done by da Silva Neto et al. (10) identified C. crescentus
38 as putatively having a Fur, a Zur, and an IscR. da Silva Neto et al. (10) studied
the putative Fur in detail and also identified genes that it regulated.
The goal of my study was to identify the iron-dependent regulators in C.
crescentus as well as other metal-dependent regulators in these same protein
families. My BLAST search returned two putative open reading frame (ORF)
matches that could putatively be from the Fur family and two from the Rrf2 family.
This differed from the work of Rodionov et al. (37) and da Silva Neto et al. (10) in
that my search found two putative Rrf2 family members instead of just one. I
hypothesize that there are two potential metal-dependent regulators in C.
crescentus in each of the two families being studied: Fur and Rrf2. Based on phylogenetic analysis, I will show that one of the two possible Rrf2 family members may be a previously unstudied protein and that the protein that da Silva
Neto et al. (10) labeled as a Fur is more closely related to known Mur proteins.
39 Methods
Culture Conditions: The C. crescentus strains used in this study were CB15
and CB2A. All strains were maintained at -80°C in stocks preserved with 10%
DMSO. Cultures were grown primarily using PYE medium (0.2% peptone, 0.1% yeast extract, 0.02% MgCl2, 0.01% CaCl2, pH 6.9). For iron-limited studies,
Hutner’s Mineral Glucose Media with slight modifications was used. Specifically,
nitriloacetic acid was eliminated, and the phosphate content was reduced by one-
half. Glucose was added to the medium after treatment with Chelex 100 to
remove any iron contamination prior to sterilization, and only ACS grade salts
were used in media. For iron insufficient cultures, glassware was acid-washed in
a 0.5 N HCl bath for at least 4 hours and was then rinsed extensively with RO
H2O. Iron sufficient cultures had iron added as FeCl3 to a final concentration of 10
µM. For metal limitation studies, 2-2’dipyridyl was added to PYE media to a final
concentration of 200 µM. For metals sufficient media, 10 µM FeCl3 was added to
PYE. All C. crescentus cultures were grown at 30°C in a shaker. For protein
preps, cultures were grown to an OD600 of 0.3-0.6, and then harvested.
Cloning: Cloning employed the use of the TOPO TA cloning kit system pCR2.1-
TOPO (Invitrogen). His-tagging of proteins was done in 2.1-TOPO and then
transferred into pRK404, a broad host range vector (42).
40 PCR: PCR reactions included the following: 0.6 ng/µL DNA template, 0.2 mM dNTPs (Promega), 0.5 µM concentration of each primer, 1X GoTaq buffer
(Promega), and 1 unit of GoTaq (Promega) per 100 µL reaction. dNTPs were kept in a 10 mM stock, and primers were kept in a 25 µM stock. PCR cycles for cloning into pCR2.1-TOPO (Invitrogen) followed manufacturer’s instructions. All other PCR, unless annealing temperature for primers required otherwise, was done using the following program: an initial step of 95°C denaturation for 3 min,
55°C annealing for 30 sec, and 72°C elongation for 2 min. This initial cycle was followed by 35 cycles of 95°C denaturation for 45 sec, 55°C annealing for 30 sec, and 72°C elongation for 2 min. After all cycles, PCR reactions were held at 10°C.
Prediction of Iron-Dependent Regulators: For an initial screen of the CB15 genome, BLAST searches employed BLASTP and the BLOSUM62 scoring matrix (2). The initial screen BLAST searches were conducted on the NCBI website using Mur (YP_766004) and RirA (YP_766387) protein sequences from
R. leguminosarum (2). Two proteins were identified in CB15 out of each of the
Fur and Rrf2 families. Members of both protein families with experimentally confirmed identities were compiled for comparison with the CB15 proteins.
Bioedit version 7.0.9.0 was utilized for pairwise comparisons which calculated percent identities and similarities using the BLOSUM62 distance matrix (22).
41 Phylogeny Estimation: For both Fur and Rrf2 families in Proteobacteria, amino
acid sequences were compiled. These included proteins with experimentally
confirmed identities and also sequences found from employing a BLAST search
using these known proteins (2). Microorganisms included in this analysis were of
the phyla Proteobacteria, and a few members of each class were selected and
used for both protein family phylogenies. The cutoff E value for being included in analysis was 8e-15 because there was an obvious difference in random match
probability with the next search result, and the BLAST search utilized whole
amino acid sequences. MUSCLE online, a program that aligns sequences while
taking into account residue specifics such as hydrophobicity and charge, was
employed to help with alignments, and the alignments produced were accessed
and edited using BioEdit version 7.0.9.0 (14, 22). Alignments took into account
known functional amino acids from Lee et al. (28) for the Fur family. Aligned
sequences were imported into ProtTest version 2.4 in order to determine the best
fit amino acid substitution model (1). For the Fur family, the model was LG+G
(weighted= 0.86); for the Rrf2 family, the LG+I+G model was used (weighted=
0.98). MrBayes version 3.1.2 was used to estimate most likely Bayesian tree topologies (24). The program was set to 2,000,000 generations with chain number set to ten, and the first 250,000 generations were discarded as burn-in.
AWTY was then used as an indicator of MrBayes completion to ensure there was
not divergence between the two runs (30). The AWTY program is used to assess
the completion of a run of Markov Chain Monte Carlo (MCMC) analyses by
42 assessing the trees produced by programs such as MrBayes (30). PhyML
version 3.0 was used to obtain best fit maximum likelihood phylogenies (21). In
PhyML, the LG model was used for both families, the proportion of invariable
sites was set to 0.0, touse simultaneousNNI was used to estimate tree
topologies, and the input tree was estimated using BioNJ. Trees were visually edited in Treegraph 2 (46).
Outer Membrane Protein Isolation: Outer membrane proteins (OMPs) were isolated using a standard protocol by Bal et al. (5). Proteins were quantified using the Pierce BCA protein quantification kit, separated on a 10% SDS-PAGE, stained with Coomassie Brilliant Blue R-250, and then photographed.
Determination of Putative Regulator Expression: CC2625 was C-terminally tagged with a poly-histidine tail using primers: 2625 F1 (5’
CTCTGACTTTCTCCGCCG 3’ Tm 55.4°C) and 2625 6-HIS (5’
TCACTAGTGATGGTGATGGTGGTGGCCTTGTATGAGTTCTGCGCC 3’ Tm
69.7 °C). The resulting product spanned the entire 431 bp coding region and also included 583 bp upstream. This was confirmed by sequencing, which indicated that the histidine residues were in-frame, and was placed into pRK404 and kept under tetracycline selection. The construct was conjugated into CB2A via strain
S17-1 and was grown in different media conditions. An empty vector was used as a control. Whole cell lysates were created by suspending cells in TE buffer
43 with 1% SDS and heating at 65°C for 15-20 min. Proteins were quantified using the Pierce BCA Protein Assay Kit (Thermo Scientific). Whole cell lysates were separated on a 15% polyacrylamide gel. PVDF membrane was pre-soaked in methanol (MeOH) for 4 min. The Mini Trans-Blot Electrophoretic Transfer Cell by
Bio-Rad was used to transfer proteins. Proteins were transferred according to
Bio-Rad manufacturer’s instructions in NET buffer (25 mM Tris, 192 mM glycine,
20% MeOH) for 30 min at a 100 V setting. Blocking, incubation with primary and secondary antibodies, and washes were all carried out according to the protocol of Smit et al. (43). The primary antibody (Sigma Monoclonal Anti-polyHistidine
Clone HIS-1Mouse ascites fluid) and the secondary antibody (GE Healthcare
ECL Plex goat-α-mouse IgG, Cy5) were both used at a 1:3000 dilution. The secondary antibody incubation was carried out with light protection, and blots were visualized using a Molecular Dynamics Typhoon imager 9400.
44 Results
Fur Family: From C. crescentus CB15, CC0057 and CC0357 were identified as potential members of the Fur family by the use of a BLAST search. As shown in
Table 2.1, CC0057 shared the greatest sequence similarity in percent identity
(58%) and percent similarity (71%) with Mur from R. leguminosarum
(YP_766004). CC0357 shared the greatest sequence similarity in percent identity
(41%) and percent similarity (54%) with Zur from Xanthomonas campestris
(YP_364737).
A phylogenetic analysis of the Fur family was performed. Both Bayesian and Maximum Likelihood means predicted the putative regulators from C. crescentus to form clades with known proteins (Figures 2.2 and 2.3). CC0357 shared homology with known Zur proteins which all grouped together into clade 1 in both analyses. CC0057, predicted by da Silva Neto et al. (10) to be a Fur protein, actually shared closer sequence homology with Murs which grouped into clade 2.
45 Table 2.1: Fur family pairwise comparison. Assessment of known metal-dependent regulators in Proteobacteria as compared with putative regulators in Caulobacter crescentus
CB15 by % identity (ID) and % similarity (sim) in amino acid sequence. Numbers were recorded using the BLOSUM62 distance matrix. No Irr homologue was found when the genome was searched using Irr from Bradyrhizobium japonicum as query. Accession numbers were taken from
NCBI. The accession numbers for the CB15 proteins were: CC0057- NP_418876, and CC0357-
NP_419176.
CC0057 CC0057 CC0357 CC0357
Accession ID (%) sim (%) ID (%) sim (%)
Fur YP_001729672 35 51 22 31
NP_231738 33 48 19 28
NP_273263 34 55 19 31
YP_001350806 32 52 17 28
Mur NP_387131 44 58 23 28
YP_766004 58 71 18 24
Zur YP_001732823 16 30 40 51
YP_364737 13 29 41 54
NP_458539 16 30 38 49
YP_001605834 16 27 39 48
46 Figure 2.2: Bayesian Fur family phylogeny. Tree includes species names as well as
NCBI accession numbers. Proteins with experimentally confirmed identities are in bold and include protein name. Clades are numbered on the right side. Arrows indicate C. crescentus
CB15 sequences. Posterior probability values are displayed on the tree.
47
Figure 2.3: Maximum Likelihood Fur family phylogeny. Tree includes species names as well as NCBI accession numbers. Proteins with experimentally confirmed identities are in bold and include protein name. Clades are numbered on the right side. Arrows indicate C. crescentus CB15 sequences. Posterior probability values are displayed on the tree.
48 Rrf2 Family: From C. crescentus CB15, CC1866 and CC2625 were identified as
potential Rrf2 family members by the use of a BLAST search. As shown in Table
2.2, CC1866 shared the greatest sequence identity in percent similarity (50%)
and percent identity (68%) with IscR from Pseudomonas aeruginosa
(YP_001346682). CC2625 shared the greatest percent sequence identity (31%)
with IscR from P. aeruginosa (YP_001346682) and shared the highest percent
similarity (49%) with RirA from Sinorhizobium meliloti (Q9R9M8). However,
CC2625 had nearly uniform percent identities and similarities with all three
known regulators of the Rrf2 family.
A phylogenetic analysis of the Rrf2 family in C. crescentus was employed
using Bayesian and Maximum Likelihood means (Figures 2.4 and 2.5). CC1866
shared sequence similarity with known IscR proteins and grouped into clade 2.
CC2625 formed a distinct clade (clade 1) with several predicted proteins from other α-Proteobacteria, including amino acid sequences from C. crescentus K31,
Asticcacaulis excentricus, Phenylobacterium zucineum, and S. meliloti. Clade 1
is a clade of proteins with an unknown function.
49 Table 2.2: Rrf2 family pairwise comparison. Assessment of known metal-dependent regulators in Proteobacteria as compared with putative regulators in Caulobacter crescentus
CB15 by % identity (ID) and % similarity (sim) in amino acid sequence. Numbers were recorded using the BLOSUM62 distance matrix. Accession numbers were taken from NCBI. The accession numbers for the CB15 proteins were: CC1866- NP_420673, and CC2625- NP_421425.
CC1866 CC1866 CC2625 CC2625
Accession ID (%) sim (%) ID (%) sim (%)
RirA Q9R9M8 21 37 30 49
NP_353235 23 42 29 48
YP_766387 23 40 30 48
IscR YP_001731461 46 67 28 44
YP_001346682 50 68 31 43
YP_001881322 46 67 28 44
YP_002425160 49 65 28 41
NsrR YP_001732944 44 52 29 47
YP_001882869 21 42 30 48
NP_458803 21 45 30 48
50
Figure 2.4: Bayesian Rrf2 family phylogeny. Tree includes species names as well as
NCBI accession numbers. Proteins with experimentally confirmed identities are in bold and include protein name. Clades are numbered on the right side. Arrows indicate C. crescentus
CB15 sequences. Posterior probability values are displayed on the tree.
51 Figure 2.5: Maximum Likelihood Rrf2 family phylogeny. Tree includes species names as well as NCBI accession numbers. Proteins with experimentally confirmed identities are in bold and include protein name. Clades are numbered on the right side. Arrows indicate C. crescentus CB15 sequences. Posterior probability values are displayed on the tree.
52 OMP Profile Data: The sequenced strain of C. crescentus is CB15, though
CB2A is often used for OMP profiles due to its lack of an RsaA layer, a surface
array found in this microbe (4, 29). In the da Silva Neto et al. (10) study that
identified CC0057 as Fur, 2,2’-dipyridyl, a powerful chelator, was used to chelate
the PYE media and simulate iron starved conditions. To simulate this condition
and compare it to minimal media, an OMP profile was performed utilizing CB2A
grown in PYE with and without 2,2’-dipyridyl and in minimal media with and
without the addition of iron (Figure 2.6). The OMP profile of cells grown in iron insufficient minimal media (lane 3) was quite different from the profile of those
grown in PYE with 2,2’-dipyridyl (lane 2). Bands present from cells grown in PYE
with 2,2’-dipyridyl (lane 2) and not in PYE with Fe3+ (lane 1) or in Hutners with
insufficient iron (lane 3) are likely to be metal receptors that are responsible for
metals other than iron.
Protein Expression: Western blots were performed in order to examine protein
expression in various media conditions. As seen in Figure 2.7, CC2625 protein
expression was dependent on metals in the medium. This protein was expressed
only when metals were removed from the medium with the addition of 2-2’-
dipyridyl (lanes 2 and 3).
53
Figure 2.6: Outer membrane protein profile of CB2A cells grown in different media. Numbers down left side are standards in kD. Arrows indicate differences in lanes 2 and
3 that are not accounted for by media components. 1: PYE with addition of 10 µM FeCl3, 2: PYE with 200 µM 2-2’dipyridyl, 3: Hutner’s mineral glucose (HMG), 4: HMG with 10 µM FeSO4, 5:
HMG with 10 µM FeCl3.
54
Figure 2.7: Western blot of HIS-tagged CC2625. Area shown is from 15 to 25 kD. All
CB2A cells were grown in PYE. 1: His-tagged CC2625, 2: His-tagged CC2625 in 2-2’-dipyridyl 3:
His-tagged CC2625 with 2-2’-dipyridyl added when cells reached OD600 of 0.4, 4: Empty vector control, 5: Empty vector control in 2-2’-dipyridyl.
55 Discussion
This analysis indicated that C. crescentus has four possible metal-
dependent regulators. Two of these are from the Fur family. Out of this family, I have demonstrated through phylogenetic analysis that there is one Mur-like protein, CC0057, and one Zur-like protein, CC0357. The other two regulators are from the Rrf2 family where I have shown through phylogenetic analysis that
CC1866 shares homology with IscR and that CC2625 is possibly a yet unstudied regulator present in several members of the class α-Proteobacteria. There are no literature references to Rrf2 family members other than IscR, RirA, or NsrR, so these were the only three members included in the phylogenies. CC2625 did not fit into a clade with any of the three known proteins from the Rrf2 family, so it is likely to represent a new member.
Recently, CC0057 from C. crescentus CB15 was proposed by da Silva
Neto et al. (10) to be the ORF encoding the Fur protein, because a strain with a
mutation in this gene was slow to grow and was sensitive to hydrogen peroxide
and organic peroxide. In that study, 2,2’-dipyridyl, a powerful chelator, was used
to chelate the PYE medium and simulate iron starvation conditions (10). 2,2’-
Dipyridyl has the ability to tightly bind to many different ions including iron, zinc
and copper (26). Figure 2.6 shows that there is a different OMP response in PYE
with 2,2’-dipyridyl than in minimal media that is simply iron deficient, so the cell
does not respond the same to both conditions. PYE with 2,2’-dipyridyl does not
56 completely mimic the response of a truly iron insufficient minimal media
condition.
C. crescentus has two forms of superoxide dismutase (SOD); one uses
iron and the other uses copper-zinc in the active site (40, 44, 45). Perhaps a chelator such as 2,2’-dipyridyl added to the medium could be interfering with
SOD or a number of other enzymes in the study by da Silva Neto et al. (10).
Chelating agents have the ability to interfere with the activity of Cu-Zn SOD by chelating the ion cofactors (27), and such interference could cause an increased level of superoxide and also an increased sensitivity to hydrogen peroxide through the Haber-Weiss reaction (Equation 3, Chapter 1). Since catalase-
peroxidase enzymes also use metals in their active sites, 2,2’-Dipyridyl could
also be interfering with the cellular catalase-peroxidase of C. crescentus (53).
This could lead to a buildup of hydrogen peroxide, feeding both Fenton and
Haber-Weiss reactions. Using PYE medium, where individual variables cannot be adequately tested, is not an effective way to study iron-dependent expression.
2,2’-Dipyridyl does not bind only iron, so using this chemical to simulate an iron insufficient condition is not reliable. I have shown that employing minimal media is a much more effective means for testing response to iron starvation.
Therefore, it is possible that C. crescentus could potentially not possess a Fur, but instead CC0057 could be a Mur as indicated by my phylogenetic analysis.
The other putative member of the Fur family that I found in the CB15 genome was CC0357. This protein is Zur-like and formed a distinct clade with
57 known Zur proteins as shown in Figures 2.2 and 2.3. Thus, I would expect this to
be a protein responsible for the zinc-dependent regulation of a number of genes
including those for uptake and use.
In the Rrf2 family, I found an IscR-like member, CC1866. This protein
formed a distinct clade with known IscR proteins as shown in Figures 2.4 and
2.5. Therefore, I expect this protein mediates the level of Fe-S clusters in the cell
by regulating genes for Fe-S biogenesis.
The second Rrf2 protein examined was CC2625, part of a distinct clade of
proteins from various α-Proteobacteria as shown in Figures 2.4 and 2.5. This
clade contained no proteins of known function, unlike RirA, IscR, and NsrR which all formed distinct clades within this family of proteins (3, 25, 48, 50). Therefore, I
conclude the clade that included CC2625 could be a novel member of this family.
In protein expression experiments, this protein was up regulated in metal limited
conditions (Figure 2.7). These data are supportive of the hypothesis that protein
CC2625 is likely a metal-dependent regulator.
Based upon these data, there are four metal-dependent regulators in C.
crescentus in the Fur and Rrf2 families. I predict out of the Fur family there is a
Zur protein and either a Mur or a Fur protein and that in the Rrf2 family, there is
an IscR and a previously unstudied regulator. This metal-regulated protein has an unknown function and is an interesting candidate for further research.
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64 CHAPTER THREE
TONB-DEPENDENT IRON UPTAKE IN C. CRESCENTUS
Introduction
The majority of what is known about high-affinity iron acquisition systems
in gram negative bacteria is from studies focused on pathogens and terrestrial
microbes, and little is known about aquatic microorganisms (2, 25). Many gram
negative bacteria employ a system that uses siderophores to tightly bind ferric
iron (Fe3+) and deliver it to TonB-dependent outer membrane receptors (30).
Although ferric-siderophore complexes are soluble, they cannot diffuse through
the outer membrane, so they are transported into the cell via these high affinity
receptors (7, 10, 20, 25, 31). Each receptor imports only one type of siderophore
and is highly specific, so bacteria generally have several receptors allowing them
to utilize more than one ferric-siderophore (2, 24).
Siderophores are chemically diverse, and the type of siderophores available can vary depending on the environment (19, 25). Thus, microorganisms need different TonB-dependent receptors based on their locations in the ecosystem (19, 25). Caulobacter crescentus is a gram negative α-
Proteobacterium that inhabits nutrient poor environments (1, 6, 11, 13, 16, 26).
Although this microorganism is often acknowledged primarily as a model for bacterial cell development and cell cycle regulation, it also has the notable ability
65 to live in extremely nutrient dilute conditions and has a large number of
annotated TonB-dependent receptors in its genome (1, 11, 12, 21).
The order Caulobacterales possesses a range of 58 to 97 annotated
TonB-dependent receptors, a much higher number than in previously studied
bacteria such as Escherichia coli which has seven, Vibrio cholerae with two,
Bradyrhizobium japonicum with three, and Pseudomonas aeruginosa with
thirteen (Figure 3.1). The C. crescentus strains CB15 and K31 are both aquatic
microorganisms as is Asticcacaulis excentricus; CB15 is found in freshwater, K31 in groundwater, and A. excentricus, a Caulobacterales member with a non-
adhesive stalk, is also found in freshwater (14, 16, 17, 26). Phenylobacterium
zucineum is a member of this order also and is an intracellular bacterium isolated
from human cells (18). Another order within α-Proteobacteria, Rhodobacterales, includes Maricaulis maris and Oceanicaulis alexandrii, both of which are morphologically similar to Caulobacteraceae and live in sea water (26, 27).
Caulobacterales have more annotated receptors than the
Rhodobacterales members M. maris and O. alexandrii which have 28 and 30 annotated receptors respectively. This is likely due to the chemistry of marine siderophores which are characteristically limited to α-hydroxycarboxylate functional groups and are typically amphiphilic, thus limiting the variety of siderophores available for uptake (19, 25). C. crescentus K31, however, being a groundwater species, most likely has access to an influx of various siderophores from other bodies of water as well as terrestrial habitats. Therefore, it would
66 V. cholerae 2
B. japonicum 3
E. coli 7
P. aeruginosa 13
M. maris 28
O. alexandrii 30
P. zucineum 58
C. crescentus CB15 66
A. excentricus 75
C. crescentus K31 97
0 20406080100 number of annotated receptors
Figure 3.1: Number of annotated TonB-dependent receptors in the genomes of several microorganisms.
67 make sense for this microbe to have the ability to utilize many different types of
siderophores that it may come across, thus it has a large number of receptors with 97 annotated in its genome. C. crescentus CB15 and A. excentricus have 66
and 75 annotated receptors respectively. These two species typically live in
freshwater and probably have access to more siderophores than marine species
and less than groundwater microorganisms. This type of relationship is reflected
in the number of receptors these species possess.
It was the goal of this study to better understand the high-affinity iron
uptake system in C. crescentus by identifying and studying the iron regulated
TonB-dependent receptors. This microorganism was grown on various
siderophores to see which it could utilize, and the outer membrane protein (OMP)
profile was examined to determine the phenotype of C. crescentus in an iron
insufficient background. I identified iron regulated receptors through reverse
transcriptase PCR (RT-PCR), RNA slot blots, and tandem mass spectrometry
(MS/MS). After identifying iron-regulated receptors, a number of gene deletion mutants were constructed, and the mutant strains were tested for their ability to grow using known ferric-siderophores as a source of iron.
68 Methods
Culture Conditions: The C. crescentus strains used in this study were CB15
and CB2A. With the exception of strain CB2A being employed for routine OMP
profiling and comparisons with CB15 growth, CB15 was the primary strain used.
All strains were maintained at -80°C in stocks preserved with 10% DMSO.
Cultures were grown in PYE medium (0.2% peptone, 0.1% yeast extract, 0.02%
MgCl2, 0.01% CaCl2, pH 6.9). For iron-limited studies, Hutner’s Mineral Glucose
Media with slight modifications was used. Specifically, nitriloacetic acid was eliminated, and the phosphate content was reduced by one-half. Glucose was added to the medium after treatment with Chelex 100 to remove any iron contamination prior to sterilization, and only ACS grade salts were used. For iron insufficient cultures, glassware was acid-washed in a 0.5 N HCl bath for at least
4 hours and was then rinsed extensively with RO H2O. Iron sufficient cultures had
iron added as FeCl3 to a final concentration of 10 µM. When siderophores were
added, they were added from a 1 mM stock of the siderophore with 0.8 mM
FeCl3. DHB (dihydroxybenzoic acid) was the only exception: it was kept in a 3
mM stock with 0.8 mM FeCl3. All siderophores were purchased from Sigma-
Aldrich with the exception of rhodotorulate which was purchased from Fisher
Scientific and enterobactin which was isolated using a protocol by Young (32). All
C. crescentus cultures were grown at 30°C in a shaker. For RNA extraction or
OMP profiles, cultures were grown to an OD600 of 0.3-0.6, and then harvested.
69 For growth curves, a small volume was removed from cultures at different time
points for an OD600 reading on a spectrophotometer (Eppendorf Biophotometer).
OMP Isolation: OMPs were typically isolated using a standard protocol by Bal et
al. (4). Proteins were quantified using a Pierce BCA protein quantification kit,
separated on a 10% SDS-PAGE, stained with Coomassie Brilliant Blue R-250,
and then photographed. OMPs used for MS/MS were isolated using the protocol
of Neugebauer et al. (21). This was due to the fact that the Bal et al. (4) protocol
utilized NP-40 which is incompatible with mass spectrometry equipment. MS/MS
analyses were done by LSU Health Sciences Center in New Orleans, LA. Before
either type of extraction, CB15 cells were treated to remove the RsaA surface array using HEPES and EGTA and following the protocol of Walker et al. (29).
Cloning and Deletions: Most cloning employed the use of the TOPO TA cloning kit system pCR2.1-TOPO (Invitrogen). Gene deletions were generated using the suicide vector pNPTS138 (a gift from Yves Brun, Indiana University) (9). The general scheme of sacB gene deletion is shown in Figure 3.2. First, two fragments of the gene were cloned (one 5’ and one 3’ of the region to be deleted) into pNPTS138 which contains kanamycin resistance and a copy of the sacB cassette. The two gene fragments were synthesized using the primers in Table
3.1 for ORFs CC0028, CC1666, and CC3336 with explanations in Figures 3.3,
3.4, and 3.5 respectively. Also, a CC3336/CC1666 double mutant was made.
70
Figure 3.2: SacB counter-selection. Deletions were constructed by inserting two sections of gene X into the suicide vector pNPTS138 via restriction digest. A large area between the two fragments of gene X was left out of the construct. The plasmid was electroporated into
CB15 and single cross-over events were selected for by kanamycin resistance and sucrose sensitivity, which indicated that the vector was inserted into the genome. A second cross-over event occurred when cells were grown without kanamycin selection, eliminating the region of the gene between the fragments originally incorporated into the vector. Second cross-over events were selected for by growth on sucrose, which is converted into levan (a polysaccharide) by the protein SacB (9). This selected for the removal of the plasmid DNA by selecting for removal of sacB and a deletion in gene X. Deletions were confirmed via PCR.
71 Table 3.1: Primers used to construct and test for deletions.
Primer name Sequence (5' to 3') Tm(°C)
0028 F2 ATGCGTAACCCGAACTTCAC 55.4
0028 R2 ACCCTTTACGACCTCGACCT 57.7
0028 F14 ATCGATGAGACGCTGAAC 52
0028 R14 AAGCTGCTCGTTGTTCTTGG 55.9
1666 F2 CTGCAGCAGCAGTTCTTCAC 56.7
1666 R2 GTGACGTCGGTCTTGTTGAA 55.2
1666 F3 TTCACGAAGTGAAGGTGA 51.4
1666 R3 TCAGTGCCTTTTCACTGAC 52.6
3336 F2 AACCGCAACACCGTCTCCG 60.7
3336 Rev AGGTGTTCTTGCCGAACAGG 58.7
3336 F6 CTCGGTTCTGTTCAACGTCA 55
3336 R6 GGCCACCTGCATGTATTTCT 55.4
3336 F3 GTGATCGACGGCGTGTAC 55.9
72
Figure 3.3: CC0028 gene deletion strategy. The black box indicates coding region and the ruler is in base pairs. The two gene fragments were synthesized in the following way: a
PCR product CC0028 F2/CC0028 R2, and a PCR product CC0028 F14/CC0028 R14 cut with
SalI/EcoRI. These two gene fragments are indicated by red line. CC0028 F/CC0028 R14 primers were used for PCR to confirm gene deletion.
73
Figure 3.4: CC1666 gene deletion strategy. The black box indicates coding region and the ruler is in base pairs. The two gene fragments were synthesized in the following way: a
PCR product CC1666 F3/CC1666 R3, and a PCR product CC1666 F2/CC1666 R2. These two gene fragments are indicated by red line. CC1666 F3/CC1666 R2 primers were used for PCR to confirm gene deletion and for sequencing of deletion.
74
Figure 3.5: CC3336 gene deletion strategy. The black box indicates coding region and the ruler is in base pairs. The two gene fragments were synthesized in the following way: a
PCR product CC3336 F2/CC3336 Rev cut with EcoRV, and a PCR product CC3336 F6/CC3336
R6. These two gene fragments are indicated by red line. CC3336 F3/CC3336 R6 primers were used for PCR to confirm gene deletion.
75 This mutant was made by taking the single gene deletion mutant CC3336 and
deleting CC1666. The deletions were constructed as described by Gonin et al.
(9). First, a single cross-over was selected for by growth on kanamycin. Then, a second cross-over was selected for through sacB selection by growth on media with 3% sucrose. The gene sacB encodes levansucrase and confers sensitivity to sucrose in gram negative bacteria (9). All deletions were confirmed by PCR using primers indicated in Figures 3.3, 3.4, and 3.5 and listed in Table 3.1.
PCR: PCR reactions included the following: 0.6 ng/µL DNA template, 0.2 mM
dNTPs (Promega), 0.5 µM concentration of each primer, 1X GoTaq buffer
(Promega), and 1 unit of GoTaq (Promega) per 100 µL reaction. dNTPs were
kept in a 10 mM stock, and primers were kept in a 25 µM stock. PCR cycles for
cloning into pCR2.1-TOPO (Invitrogen) followed manufacturer’s instructions. All
other PCR, unless annealing temperature for primers required otherwise, was
done using the following program: an initial step of 95°C denaturation for 3 min,
55°C annealing for 30 sec, and 72°C elongation for 2 min. This initial cycle was
followed by 35 cycles of 95°C denaturation for 45 sec, 55°C annealing for 30 sec,
and 72°C elongation for 2 min. After all cycles, PCR reactions were held at 10°C.
RNA Extraction and RT-PCR: RNA was extracted from cells using Trizol
reagent (Invitrogen) and the protocol used by Hottes et al. (11). Once extracted,
RNA was run on a 1% agarose gel stained with ethidium bromide to check for
76 integrity, and was also quantified using UV spectroscopy. Reverse transcriptase
PCR (RT-PCR) was done using the Superscript III One-Step RT-PCR system with Platinum Taq (Invitrogen), and cycles were programmed according to manufacturer’s instructions. The primers that were used are listed in Table 3.2 and spanned an area of ~500 bp with a forward primer ~200 bp into the annotated coding region and a reverse primer ~700 into the annotated coding region. RNA for each media condition was extracted from a single culture. For each 50 µL reaction, 1 µg of RNA was included, and resulting cDNA was run on a 1.5% agarose gel and stained with ethidium bromide for visualization.
Slot Blots: Slot blots were done using the protocol of Page et al. (23). RNA extracted from cells grown in different conditions was arrayed to Hybond-N+ membrane (Amersham) using a slot blot manifold. Blots were then UV cross- linked in a Bio-Rad GS Gene Linker UV Chamber at the 150 mJoule setting.
Each growth condition had four different amounts of RNA immobilized on the membrane. This process was repeated at least three times using RNA extracted separately. A PCR product for the ORF (open reading frame) of interest was obtained using the same primer pairs used in RT-PCR, listed in Table 3.2.
Labeling of PCR products followed the protocol of Feinberg et al. (8). Twenty microliters of PCR product was purified via a PCR Purification Kit (Qiagen) and boiled for 10 min. Then 25 µL of labeling buffer (1 M HEPES pH 6.6, 0.25 M Tris,
25 µM MgCl2, 0.36% β-mercaptoethanol, 0.42 µg/µL Random Primers from
77 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF.
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
0028 F2 ATGCGTAACCCGAACTTCAC 55.4 485
0028 R2 ACCCTTTACGACCTCGACCT 57.7
0139 F2 GATCGCCGGTTCGAGCCATG 61.1 499
0139 R2 CCGTATTGTCGCGAACGCCG 61.5
0171 F2 AGATCCAGAGCTCGCTGAAC 56.8 476
0171 R2 CCACAGATCTTCGTCCAGGT 56.5
0185 F2 CCGACCCTGCAGTTCTATTC 55.3 488
0185 R2 AGGTCTTCACTGGGCTTGAA 56.3
0210 F2 GCAAGCCCTACAACGACTTC 56.2 474
0210 R2 CCGTGATGATGATGTTTTCG 51.8
0214 F2 GGTCATCGTCACCGCCGCAC 63.7 520
0214 R2 GCTCTCGCGCTTCAGGGCGT 65.3
78 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
0442 F2 GCTTCAACTACGGCTTCGAC 56.1 530
0442 R2 GAAGTCGCCCTGGATGTAGA 56.3
0446 F2 GCCCAGCAACTATCTGAAGG 55.7 540
0446 R2 CTCGACTTGTAGGGCTCCAG 57.0
0454 F2 GTGCAGAACCAGTCGAACAA 55.6 535
0454 R2 TGTTGCTCTTGAACGAGGTG 55.3
0539 F2 GCCAAGCATAACGAAGAAGC 54.6 524
0539 R2 ATCGCATAGACCGAGGTCAG 56.3
0562 F2 CCGAACCTTCGTGATCTTTC 53.4 474
0562 R2 ACCACTTCGGAAAAGCCTTC 55.5
0571 F2 CCGACCTCGCTCTACAATTC 55.2 471
0571 R2 GTAGGCTTGGGTGTTGTCGT 57.5
79 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
0579 F2 TCCCAGCTTCAACGATCTCT 55.7 498
0579 R2 GTCAGGCCTGCATCATAGGT 57.0
0663 F2 GTCCAGATCCTGCCGTTCT 56.9 479
0663 R2 GTGTTGATCGGATCCTCCAG 55.1
0722 F2 AGAACCTGTCGCTGTTCGTC 57.3 476
0722 R2 CTCCACCTTCATCTGCCAGT 56.9
0815 F2 AAGGGTCCGACGACCTATCT 57.3 513
0815 R2 ATTGACCTTGGTCTCGTTGC 55.5
0970 F2 AAGAACGGTTTCACCGTCAC 55.5 526
0970 R2 AGAACGAGTTCACCCACCAG 56.9
0983 F2 TACTACCACGCGCTGTTCAC 57.3 504
0983 R2 GCTGATGCGATAGACGTTGA 54.9
80 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
0991 F2 GTTCCATCGCTGAAGGTCTC 55.5 516
0991 R2 GTTGTAGTCGGCGATGAAGC 56.2
0995 F2 ACACCAGCACCGCCTATATC 56.9 481
0995 R2 CCGAACTCGAACTTGGTCTC 55.3
0999 F2 CACCTCGAAGATGGTCCAGT 56.5 502
0999 R2 GGGCAGCAGGTTGTTGTACT 57.9
1093 F2 CCAGTTCAGCTCGATCTTCC 55.3 481
1093 R2 CAGATAGTTGGCGCAGTTCA 55.2
1099 F2 ACTTCTACACCGGCGACATC 56.9 506
1099 R2 GTCCTTGCGATAGTGGAAGC 55.8
1113 F2 CCAGGTTCGTCTGTTCTGGT 56.9 525
1113 R2 GTGCTTTCGTTGGTCTTGGT 55.9
81 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
1131 F2 ATACCAAGCTCGACGACCAC 56.9 545
1131 R2 GATATCCGCGCGATTGTAGT 54.8
1136 F2 AACTCGTCCATTTCCGTCAG 54.9 550
1136 R2 TCAGGTTGGCCTTGATCTG 54.7
1138 F2 AATGATCCGTTCGGTGGTT 54.4 523
1138 R2 CGGAGCTGTTCTCAAGATCG 55.4
1142 F2 TGCAGCTCAATGAGAACGAC 55.4 478
1142 R2 CAGTACAGCGTGCCCAGATA 56.8
1145 F2 TCGATCTCAATGTCCTGACG 54.1 509
1145 R2 GAGCTTGTAGCGCTGGATGT 57.6
1517 F2 GCTTCCAGGTGCTCAAGG 56.2 461
1517 R2 GTAGGTCGTGGTCAGCTGGT 59.0
82 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
1623 F2 GATGCGGTTGAGGAGATCAT 54.6 517
1623 R2 AGTTGGCCATCGTCGAATAG 54.8
1666 F2 CTGCAGCAGCAGTTCTTCAC 56.7 455
1666 R2 GTGACGTCGGTCTTGTTGAA 55.2
1750 F2 GCAACCGATCGAGAAGGTC 55.7 490
1750 R2 ACCGTCGGTGGTGTAATAGC 57.0
1754 F2 CCGTCGAGTTCGACCAGTAT 56.3 460
1754 R2 TTCGGCTGGTACTCGAAGAT 55.7
1778 F2 GGGATCATCAACGTCGTCTC 55.2 486
1778 R2 GAAGGACGACCATTCCTTGA 54.4
1781 F2 ATGGTGTTCCCACTGGACAT 56.4 457
1781 R2 GCAGGCTGTACTGACCCATC 57.9
83 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
1791 F2 TATCTGGAGACCCTGCCATC 56.0 492
1791 R2 AGTCGTCCGTACGCTGAGAT 57.9
1809 F2 CGTGGGACATCAATGACAAG 53.8 466
1809 R2 TTCGAATCCAGGTGATAGGG 53.8
2149 F2 TCGACGCTGTCGTCATTATC 54.6 543
2149 R2 CCTTCCACTTGTCGCTCTTC 55.7
2194 F2 CTGGCGTTTCGGTGCGCA 62.1 586
2194 R2 TGGTCGACCGTCAGGCGG 62.8
2287 F2 ATGACCCGACCTATCTGCTG 56.4 518
2287 R2 GCGACCGAAATACTTCTCCA 54.7
2398 F2 CTCTTAACATCGGCCAGCTC 55.6 485
2398 R2 GCTGAACGTCTGCTTGTTGA 55.8
84 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
2660 F2 CCCAGACGCTGTATTTCGAT 54.8 503
2660 R2 GACCGTGTAGCTGAGGAAGC 57.8
2664 F2 GGTGCTGGAGTTCATCAAGC 56.4 550
2664 R2 ATAGTTCACGTACGGGCTGG 56.8
2709 F2 GGGACGATCAATCTGGAGAA 54.0 500
2709 R2 CCGATGAAGGGATTGAAAAA 50.7
2804 F2 TTCCTCTACAGCGACCGTTT 56.1 547
2804 R2 CAGTGGTCTGGTTGGTGTTG 56.2
2819 F2 ACGGCAGCAACTCGCTGCTG 63.5 512
2819 R2 GGCGCGCAGCATCAGGTCGT 65.8
2820 F2 GTCGAAGAAATCGTCGTCAC 53.5 550
2820 R2 GGGCTTCACGCTTGTTGTAG 56.5
85 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
2832 F2 GCTACGAGACCATCGGCTAC 57.3 479
2832 R2 GAAGTTCGAGTTGGCGTAGG 55.6
2924 F2 GACTTCTCGGCCACCTATCA 56.3 491
2924 R2 ACGCCCGAATAGGTCTTCTT 56.0
2928 F2 GCGTCGCCGACGTGCTGCAG 66.6 500
2928 R2 GCCCAGATCGGCCGAGAAGC 63.3
3001 F2 GCGCCGTCGACATCTATATT 54.8 532
3001 R2 ATTTGAACCGGCTTGTTGTC 53.9
3013 F2 ATCACGGCCTTCAATACGAC 55.0 533
3013 R2 TGGACGTTCTTTTCCTGGAC 54.9
3127 F2 GTGTGTCGTTCACCAGTTCG 56.2 478
3127 R2 GTGAAGTCGTTGGTCCGATT 55.1
86 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
3146 F2 GCTGCAGTTCACCTCAGACA 57.5 480
3146 R2 AGCGAGTCGGTCTTCACATT 56.4
3147 F2 TCGGAATTTGAGACCTACGG 54.2 533
3147 R2 TTGCTGAACGTCTTGGTGAG 55.3
3161 F2 CGTGACCAACACCTTCAATG 54.2 529
3161 R2 GACGTTGCGGTTGTAGTCCT 57.4
3194 F2 GTCTCTTCTGGCGAAGGTTG 55.7 460
3194 R2 ACCACGACTTCTTCGACCTG 56.8
3336 F6 CTCGGTTCTGTTCAACGTCA 55.0 514
3336 R6 GGCCACCTGCATGTATTTCT 55.4
3436 F2 AGGGTCGCTATTTCGCCTAT 55.9 466
3436 R2 TGAGTGCGGTGATAGTCGTC 56.6
87 Table 3.2: Primer pairs used for RT-PCR and for RNA slot blots for each
ORF (continued).
Product size
Primer name Sequence (5' to 3') Tm(°C) (bp)
3461 F2 CAAGGTCGACAAGAACGTCA 55.0 454
3461 R2 CAGTTCCTGGGTGAACTGCT 57.3
3500 F2 TCAATGGCTCGAACATCAAC 53.2 522
3500 R2 GGATGGCGAAGCTTTGATAG 53.6
88 Invitrogen, 0.1 mM of each of the dNTPs: dGTP, dTTP, and dATP which were
purchased from Promega) was added. Then, 1 µL of Klenow (Promega) was
added along with 1 µL of [α-32P]-dCTP for labeling which was carried out at 37°C
for 1 hr. The probe was then check to ensure that the label was incorporated.
This was done by making sure the probe had a reading of at least 20,000 cpm on
a Geiger Counter. Then probe was purified using a sephadex G-50 column and was denatured at 95°C for 10 min. Blots were pre-hybridized with 120 µg/mL sheared herring sperm DNA (Sigma), 0.1% SDS, and 3X SSC at 65°C for at least 6 hours. The probe was added to the pre-hybridization buffer and hybridized at 65°C overnight. The blot was washed two times for 20 min at 65°C with 1X blot wash (3X SSC, 5 mM EDTA, 0.02% SDS) and two more times for 20 min at 65°C with 0.1X blot wash. Blots were developed using a phosphor imager and visualized using a Molecular Dynamics Typhoon imager 9400. Hybridization quantification was done using Image Quant software (GE Healthcare). Blots were stripped and hybridized with a 16S rRNA PCR product for normalization.
89 Results
The sequenced strain of C. crescentus is CB15, though CB2A is
frequently employed for OMP profiles due to its lack of an RsaA layer, a surface
array often found in this microbe (3, 22). To identify putative iron-regulated TonB-
dependent receptor proteins in C. crescentus, CB2A cells were grown in iron
sufficient and insufficient conditions. Cells grown in insufficient iron expressed
five protein bands that were not present in iron sufficient conditions as shown by
OMP profile (Figure 3.6A). These five bands were hypothesized to be iron-
regulated TonB-dependent receptors for C. crescentus and suggest that C. crescentus has the ability to utilize at least five different ferric-siderophores. The five bands were excised and sent off for MS/MS analysis, and results can be seen in Figure 3.6B. The proteins identified were CC1754, CC1666, CC0028,
CC3146, CC3147, CC2194, CC2928, and CC1750.
The ability of two C. crescentus strains to grow on FeCl3 was tested, and both strains had robust growth with the addition of 10 µM FeCl3 (Figure 3.7).
Various siderophores were added to minimal media to test the ability of both
strains to utilize different iron chelators for growth. Both strains were able to
utilize ferric-rhodotorulate (Figure 3.8), ferric-citrate (Figure 3.9), ferric-DHB
(Figure 3.10), ferric-ferrichrome (Figure 3.11), and ferric-enterobactin (Figure
3.12). There was a difference in the ability of the strains to utilize ferric-
ferrichrome. CB2A would grow on 1 µM ferrichrome while a larger concentration
90
Figure 3.6: Outer membrane profile and MS/MS data for CB2A. A: Outer
membrane profiles (OMPs) of CB2A cells grown in minimal glucose media with 10 µM FeCl3
(+Fe) and in minimal glucose media with insufficient iron (-Fe). The numbers on the side represent size in kD. B: The bands only present when grown in insufficient iron are putative iron- regulated TonB-dependent receptors and were identified through MS/MS.
91 A.
B.
Figure 3.7: Growth at OD600 of two C. crescentus strains on FeCl3. Strains
CB2A and CB15 were grown in Hutner’s minimal media with the addition of FeCl3, and optical density was measured. Closed shapes represent CB2A, and open shapes represent CB15.
Circles represent the addition of FeCl3. Triangles indicate no FeCl3 was added. A: 1 µM FeCl3 added for circle shapes, B: 10 µM FeCl3 added for circle shapes. Note that the y-axis for the two graphs is different. B has double the coverage of y values of A.
92 A.
B.
Figure 3.8: Growth at OD600 of two C. crescentus strains on ferric- rhodotorulate. Strains CB2A and CB15 were grown in Hutner’s minimal media with the addition of ferric-rhodotorulate, and optical density was measured. A: 1 µM rhodotorulate, B: 10
µM rhodotorulate. Closed circles represent CB2A, and open circles represent CB15.
93 A.
B.
Figure 3.9: Growth at OD600 of two C. crescentus strains on ferric-citrate.
Strains CB2A and CB15 were grown in Hutner’s minimal media with the addition of ferric-citrate, and optical density was measured. A: 1 µM citrate, B: 10 µM citrate. Closed circles represent
CB2A, and open circles represent CB15.
94 A.
B.
Figure 3.10: Growth at OD600 of two C. crescentus strains on ferric-DHB.
Strains CB2A and CB15 were grown in Hutner’s minimal media with the addition of ferric-DHB, and optical density was measured. A: 1 µM DHB, B: 10 µM DHB. Closed circles represent CB2A, and open circles represent CB15.
95 A.
B.
Figure 3.11: Growth at OD600 of two C. crescentus strains on ferric- ferrichrome. Strains CB2A and CB15 were grown in Hutner’s minimal media with the addition of ferric-ferrichrome, and optical density was measured. A: 1 µM ferrichrome, B: 10 µM ferrichrome. Closed circles represent CB2A, and open circles represent CB15.
96 A.
B.
Figure 3.12: Growth at OD600 of two C. crescentus strains on ferric- enterobactin. Strains CB2A and CB15 were grown in Hutner’s minimal media with the addition of ferric-enterobactin, and optical density was measured. A: 1 µM enterobactin, B: 10 µM enterobactin. Closed circles represent CB2A, and open circles represent CB15.
97 (10 µM) was needed in order for CB15 to grow (Figure 3.11). Neither strain would grow on the ferric-chelate ferric-pyoverdines.
TonB-dependent receptors with putative iron regulation were identified with RT-PCR. RNA was isolated and was used to screen the 66 TonB-dependent receptors in C. crescentus. RT-PCR identified seven ORFs that putatively had higher transcript level in insufficient iron over sufficient iron (10 µM FeCl3)
conditions as shown in Figure 3.13. The seven identified were CC0028, CC3336,
CC2924, CC2194, CC1623, CC2832, and CC0454.
RNA slot blots were performed to look at the expression levels of genes
identified from RT-PCR. Slot blots identified two ORFs that had higher transcript
level in iron insufficient than in iron sufficient conditions as shown in Table 3.3.
These ORFs, CC0028 and CC3336, both showed about two fold transcript level
in iron insufficient over iron sufficient conditions. RNA blots were attempted for
various other ORFs, but reproducible results were not attained. From three
experiments, RT-PCR, RNA blots, and MS/MS, nine genes were identified as
putative iron-regulated TonB-dependent receptors (Table 3.4).
Gene deletions were constructed in strain CB15 in ORFs CC3336,
CC0028, CC1666, and also a double mutant of CC3336/CC1666. The PCR
products spanning these deletions are shown in Figure 3.14. The CC1666
deletion was confirmed by sequencing and the deletion details are shown in
Figure 3.15. A large region was deleted, and the surrounding DNA sequence was
98
Figure 3.13: Reverse transcriptase PCR (RT-PCR) to identify iron regulated
TonB-dependent receptors. RT-PCR was performed using primers for all 66 putative
TonB-dependent receptors. The first seven shown have possible up regulation in insufficient iron.
The bottom two, CC1754 and CC3147, were identified as having iron-dependent regulation in a different experiment, so were included in this figure. “+” indicates that 10 µM FeCl3 was added to the minimal medium.
99 Table 3.3: Slot blots to confirm the iron regulation of TonB-dependent receptors. Included are six ORFs that RT-PCR indicated were putatively iron-regulated
(CC0028, CC3336, CC1623, CC0454, CC2194, and CC2832) and three additional ORFs for negative controls (CC2820, CC0139, and CC1517). Fold difference refers to transcript levels for cells grown in insufficient iron over sufficient iron. The number of replicates is in parenthesis.
CC3336 and CC0028 had about 2.5 fold and 2.0 fold increases respectively in insufficient iron over sufficient iron (10 µM FeCl3). The other four ORFs with possible iron regulation did not show a major difference in transcript concentration over the two conditions. One of the negative controls, CC0139, and one of the ORFs identified through RT-PCR but that showed no regulation in RNA blots, CC2194, were both indicated by da Silva Neto et al. (5) to be Fur regulated.
ORF Fold Difference
CC0028 2.45 ± 0.52 (8)
CC3336 2.05 ± 0.28 (11)
CC1623 1.22 ± 0.2 (12)
CC0454 1.16 ± 0.09 (11)
CC2194 0.87 ± 0.12 (21)
CC2832 0.75 ± 0.14 (10)
CC2820 0.87 ± 0.25 (11)
CC0139 1.12 ± 0.07 (12)
CC1517 1.15 ± 0.15 (10)
100 Table 3.4: Summary of findings from three experiments: RT-PCR, slot blots, and
MS/MS. ORFs included were identified as iron regulated in slot blots and/or MS/MS. “+” indicates up regulation in insufficient iron conditions over sufficient iron (10 µM FeCl3). “–“ indicates the opposite of “+”,“nd” indicates no data, and “x” indicates not tested. “+/-“ indicates that the levels remained the same for both conditions.
ORF RT-PCR Slot Blots MS/MS
CC0028 + + +
CC3336 + + nd
CC2194 + - +
CC1666 x nd +
CC3146 - x +
CC1750 x nd +
CC1754 +/- nd +
CC2928 x x +
CC3147 +/- x +
101
Figure 3.14: Deletion confirmations through PCR. For each picture, (1) is a PCR product from CB15 and (2) is a PCR product from a CB15 deletion mutant. The numbers on the side indicate approximate size in base pairs. A: CC0028, B: CC1666, C: CC3336, D: CC3336 mutant was further mutated with a CC1666 deletion. The PCR product confirms the 1666 deletion in this double mutant.
102
Figure 3.15: Map of 1666 deletion in CB15. A: the intact gene is seen. CC1666 primers F3 and R2 were used to sequence the deletion. The white box indicates coding sequence
(CDS), B: the deleted gene. Lettered DNA fragments from A are seen in a different arrangement in the deletion mutant. A large portion (1441 bp) of CC1666 was deleted, and regions A, B, C, and D remain but are in a different order. Fragment D has been reversed. In intact gene CC1666,
Fragment A begins at the start of the CDS and is 89 bp in size. Fragment B begins 1530 bp upstream of the CDS and ends 1851 bp downstream. Fragment C begins 1522 bp upstream of the CDS and ends 2032 bp downstream. Fragment D begins 1843 bp upstream of the CDS and ends 2089 bp downstream.
103 rearranged. Fragment A was 89 bp at the beginning of the coding sequence.
Between fragment A and the remaining fragments (B, C, and D), a large region of
DNA (1441 bp) was deleted. Fragments A, B, C, and D are the only regions of the CC1666 gene present in the deletion mutant. Fragments B, C, and D overlap one another in the intact gene and are 320, 510, and 246 bp in length respectively. Fragment D is reversed in the deletion mutant. All CB15 mutants were grown in minimal media with different siderophores added, and none had growth that differed from wild type. Because of a recent study indicating that
CC3336 is also carbon regulated (15), the CB15 CC3336 deletion strain was also grown under iron and carbon starvation, but no growth phenotype was seen
(Figure 3.16).
104 A. B.
C. D.
Figure 3.16: Growth of CB15 CC3336 deletion mutant on various concentrations of iron and carbon. Error bars represent the standard error from three replicates done side-by-side and taken from the same original media stock. Optical density was measured over time for cultures of CB15 (closed circles) and CB15 with a deletion in CC3336
(open circles). A: 0.5 µM ferric-rhodotorulate, 0.01% glucose, B: 1 µM ferric-rhodotorulate, 0.01% glucose, C: 0.5 µM ferric-rhodotorulate, 0.1% glucose, D: 1 µM ferric-rhodotorulate, 0.1% glucose. Note that C and D have a ~4 fold greater y-axis scale than A and B.
105 Discussion
C. crescentus has the ability to grow on ferric-ferrichrome, ferric-DHB,
ferric-citrate, ferric-rhodotorulate, and ferric-enterobactin, but not ferric- pyoverdines (Figures 3.8-3.12). This differs from a previous study done by
Neugebauer et al. (21) which showed that C. crescentus could not grow on ferric- ferrichrome, ferric-citrate, and ferric-DHB. That study chelated the entire medium with 2-2’-dipyridyl or EDDA and used a different strain than I employed (21).
Differences between that study and mine could be due to strain variation or to differences in media components. It is known that each TonB-dependent receptor imports only one type of siderophore and is highly specific (2, 24).
Therefore, through siderophore growth data (Figures 3.8-3.12) and OMP profile
(Figure 3.6A) to pinpoint proteins present only in iron insufficient conditions, I conclude that C. crescentus has at least five iron-regulated TonB-dependent receptors.
Recently, da Silva Neto et al. identified four receptors in C. crescentus as being regulated by a protein they identified as Fur in response to iron, and they predicted four more receptors to have a predicted weak Fur binding site.
However, this study used PYE with 2-2’-dipyridyl to simulate iron insufficient conditions (5). In Figure 2.6, I show that PYE with 2,2’-dipyridyl does not completely mimic the response of a truly iron insufficient minimal media condition. 2,2’-Dipyridyl has the ability to tightly bind to many different metal ions and is not iron specific, so the receptors da Silva Neto et al. (5) identified as iron
106 specific have the potential to be receptors for a variety of other metals. Through
RT-PCR, RNA blots, and MS/MS, I have limited the list of iron-dependent receptors to nine (Table 3.4). When compared to the data of da Silva Neto (5), the list of potential metal-dependent and iron-dependent TonB-dependent receptors expands to a total of 13 ORFs.
Two of the ORFs da Silva Neto et al. (5) identified as “iron” regulated,
CC0139 and CC2194, were shown by my RNA blot data not to have iron- dependent regulation. This inconsistency could be due to the da Silva Neto et al.
(5) study chelating other metals out of the media in addition to iron. If this is the case, CC0139 and CC2194 are likely to be receptors for some other metal.
CC0028 and CC2928 were identified by various combinations of my three experiments to be up regulated in iron-limited conditions. This was also the case when these were studied by da Silva Neto et al. (5). In my study, a CC0028 mutant displayed no apparent phenotype when grown on various siderophores, though it is likely that this receptor recognizes a ligand not tested in my study.
Two other ORFs that I identified as being iron regulated and for which deletions were constructed were CC3336 and CC1666. CC1666 was identified only through MS/MS, so it is only putatively iron regulated, but CC3336 was identified through RNA blots as being consistently up regulated in iron starvation.
Single gene knockout strains and a double gene knockout displayed no apparent phenotype when grown on various siderophores. A recent study done by Landt et al. (15) identified CC3336 as being post-transcripionally activated during carbon
107 starvation by a sRNA. This, combined with my data, indicate that CC3336 is under the regulation of two different nutrients: iron and carbon. Therefore, I tested the CC3336 mutant for growth under iron and carbon starvation but saw no phenotype (Figure 3.16). This could indicate that this gene may have a very complex regulation, and the right conditions have yet to be found in order to observe a phenotype.
The lack of phenotype in any receptor deletion mutants could perhaps be due to redundancy in the genome of C. crescentus. In other words, there may be more than one gene to perform a specific function, and this has been reported to occur in C. crescentus in genes involved in holdfast synthesis in stalked cells
(28). It is possible that there may also be some redundancy in iron uptake genes.
When a deletion is made, if there is more than one gene to take in ferric- ferrichrome, for example, a second gene may be able to produce enough ferric- ferrichrome receptors for the cell to obtain adequate iron levels. Another possibility is that the receptors that have been deleted are receptors for siderophores that have not yet been tested.
Overall, through this study, a number of siderophores that C. crescentus has the ability to utilize have been identified. Also, nine putative iron-regulated
TonB-dependent receptors have been found, with CC0028 and CC3336 being confirmed through RNA blots. Further studies will need to be done in order to confirm the identification of the others and to clarify the level of iron regulation.
More single and multiple gene deletions will need to be constructed to test
108 growth on siderophores, and it would also be helpful to investigate siderophores that have been previously untested. A more extensive series of deletions and growth tests will help to match up receptors with the type of ferric-siderophore they bring into the cell.
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113 CHAPTER FOUR
SUMMARY AND FUTURE DIRECTIONS
Introduction
Caulobacter crescentus is known as a model for bacterial development
and cell cycle regulation. Because of this, there are many genetic tools available
for the study of this organism. C. crescentus is also quite interesting because it is
able to live in oligotrophic conditions and is even found in commercially sold
bottled water (1, 10, 11, 17). Due to C. crescentus’s ability to grow in such
extreme oligotrophic conditions and the availability of a large number of tools for
the study of this microbe, it is an easy step to propose this microorganism as a model for life in oligotrophic freshwater conditions. Thus, my research focused on
C. crescentus and its ability to obtain enough iron to survive in its habitat.
Due to iron insolubility and toxicity in aerobic environments, bacteria have
developed systems to tightly regulate iron uptake, storage, and use. An
assortment of proteins mediate this response to iron and other required metals in
order to keep each at an adequate level for cellular homeostasis (6, 8, 12, 22).
One of the ways iron-dependent regulation takes place is at the level of iron
uptake. Although little is known about high-affinity iron acquisition systems in
aquatic species, many gram negative bacteria employ a system that uses
siderophores to tightly bind ferric iron (Fe3+) and deliver it to high affinity TonB-
dependent outer membrane receptors (2, 22, 27). Each receptor imports only
114 one type of siderophore and is highly specific, so bacteria generally have several receptors to be able to utilize numerous siderophores (2, 21).
The goal of this study was put together a model of iron-dependent uptake and metal-dependent regulation in C. crescentus. This freshwater living microbe, as well as the rest of the order Caulobacterales, has a large number of annotated
TonB-dependent receptors as compared with other microorganisms (Figure 3.1).
C. crescentus also has been studied very little with respect to metal-dependent regulators. Due to the numerous laboratory tools available for the study of this microorganism and its ability to live in extremely oligotrophic conditions, C. crescentus makes a good model for the study of metal-dependent regulation and iron uptake in freshwater microorganisms.
Metal-dependent regulators
Phylogenetic analysis showed that C. crescentus has four possible metal- dependent regulators. Two of these are from the Fur family. Out of this family, my data indicate that there is one Mur-like protein, CC0057, and one Zur-like protein,
CC0357 (Figures 2.2 and 2.3). The other two regulators are from the Rrf2 family where I have shown that CC1866 shares homology with IscR and that CC2625 is a possibly yet unstudied regulator present in several members of the α-
Proteobacteria (Figures 2.4 and 2.5).
Previous work by da Silva Neto (3) suggested that CC0057 in C. crescentus CB15 encodes a Fur protein. They performed this study by
115 constructing a strain lacking this gene (3). The resulting mutant strain was slow
to grow in the presence of hydrogen peroxide and organic peroxide. da Silva
Neto et al. (3) also identified several genes, including TonB-dependent receptors, that were “Fur” regulated. My study confirmed the iron regulation of a few of these. While CC0057 may actually be Fur, the conditions that resulted in derepression are still unclear (3). Iron insufficient conditions were obtained through the addition of 2,2’-dipyridyl to PYE, which is a very ineffective means for removing iron (3). This data would have been much more convincing had the study used minimal media. These two different conditions promote very different
phenotypes (Figure 2.6). For example, two of the four “iron” regulated ORFs
identified by da Silva Neto et al. (3) showed no increase in transcript levels in my
minimal media study (CC0139 and CC2194) (Table 3.3). It is possible that the results of this study were flawed, and CC0057 could be a Mur as indicated by my phylogenetic analysis (3). Manganese and iron-dependent regulation are often interchangeable, and manganese has even been seen to interfere with iron homeostasis (20). Further studies on this gene will first need to clarify if it is acting as a Fur or a Mur, repeating some of the key experiments done by da
Silva Neto et al. (3) but in minimal instead of complex medium. Then the results
would be more likely to have been obtained due to iron limitation and not some
other component of the medium.
In the Rrf2 family, one of the members I found was the protein CC2625.
When examined phylogenetically, this protein was part of a distinct clade of
116 proteins with members from α-Proteobacteria. This C. crescentus novel
regulator, CC2625, was up regulated in metal limited conditions (Figure 2.7). The
Rrf2 family members studied so far all rely on Fe-S clusters. IscR is an Fe-S
dependent regulator, and RirA is also thought to be regulated by Fe-S (23, 25).
NsrR requires an Fe-S cluster in order to bind to DNA in response to nitric oxide
(23, 25). Because of this, it seems likely that this novel member of the Rrf2 family
(CC2625) is also Fe-S dependent. Thus, I examined the Fe-S binding sites in the
Rrf2 family of proteins. This is shown in Figure 4.1 where the C-terminal cysteine
residues coordinating Fe-S cluster binding in IscR from E. coli are highlighted
(23) and compared to cysteines in the C-terminus of the two C. crescentus putative Rrf2 family members and to known proteins: RirA and NsrR. While the putative C. crescentus IscR (CC1866), S. meliloti RirA, and E. coli NsrR are very similar to IscR from E. coli in putative Fe-S binding site, CC2625 is quite different. CC2625 has cysteines in the same general area as the other proteins but has a different number of cysteines that form a quite different motif from the others (Figure 4.1). This C-terminal CXXC motif seen in CC2625 is present in a variety of known metal-binding proteins and in various enzymes such as thioredoxins (4, 9). CXXC is used as a redox switch for the reduction of disulfide bonds and other forms of oxidized cysteines when it is employed by thioredoxins
(9). This motif is also a known zinc binding domain both for the Zur protein and for Fur (14, 16). Also, CXXC binds copper ions and is present in CopZ, a copper transport protein (4, 30). Additionally, PerR, a redox-sensing transcriptional
117
Figure 4.1: The [2Fe-2S] coordinating site of IscR as comparison for other proteins. The top protein in this figure is IscR from E. coli (23). This protein has a known Fe-S binding site, and the important residues are highlighted in red. Corresponding cysteines in various other proteins are highlighted in red and numbers correspond to their place in the amino acid sequence. There is a blue box around the CXXC sequence present in CC2625, a putative novel regulator of the Rrf2 family.
118 regulator, contains this motif which is thought to be a metal binding site for
peroxide sensing (26, 29). In general, the CXXC motif is seen in certain enzymes
such as thioredoxins and in metal binding proteins where it binds various metals
including zinc, copper, mercury, and cadmium (4, 9). The metal to which CC2625
responds is still a mystery, but this CXXC domain indicates possible metal-
binding and could be a “switch” for the regulatory action of this protein on target genes.
In future studies, the strain with His-tagged CC2625 could be grown in various metal conditions for Western blots in order to gain a better understanding of how CC2625 is regulated, and, therefore, what metal is most likely its counterpart in gene regulation. Perhaps this protein is a new type of iron- dependent regulator. If my phylogeny based assessment is correct, there has been no iron-dependent regulator identified in C. crescentus. Perhaps iron- dependent regulation is mediated via this new member of the Rrf2 family.
Because there is a strain with CC2625 His-tagged, in the future, it would also be beneficial to purify this protein. Then, when genes regulated by this protein are identified, binding can be examined, especially in the context of CC2625’s counterpart metal. This would give us a better idea of the scope of CC2625 as a regulator and determine the function of this protein. Mutations in the CXXC site of this protein to see if this binding still occurs would also be beneficial to this study, because this would show if this motif is important to the function of protein
CC2625.
119 Iron-regulated TonB-dependent receptors
It is known that each TonB-dependent receptor imports only one type of
siderophore and is highly specific (2, 21). Therefore, because I found that C. crescentus can grow on at least five different siderophores, there are at least five iron receptors in this microorganism (Figures 3.8-3.12). However, it is likely that there are probably more than five iron-regulated receptors in this microbe. It would be beneficial for C. crescentus to be able to utilize as many siderophores as possible depending on the environment that it finds itself in, and I tested only a relative few of the siderophores known to exist. There are 66 annotated TonB- dependent receptors in this microorganism. To identify those that were iron- regulated, three different experimental approaches were used. In all, nine possible genes were identified (Table 3.4).
The regulation on some of these genes may be complex. Three of the genes I identified (CC0028, CC2194, and CC2928) were confirmed by da Silva
Neto et al. (3) to be metal regulated. That study pinpointed CC0139 and CC2194 as being iron regulated while my RNA blot data (Table 3.3) indicated no iron regulation (3). CC0139 and CC2194 are likely to be regulated by a metal besides iron and were picked up by the da Silva Neto (3) study due to their use of 2-2’- dipyridyl. In a recent study done by Landt et al. (13), CC3161 was found to be carbon regulated, and in the study done by da Silva Neto (3) this protein was predicted to have a weak Fur binding site. However, da Silva Neto et al. (3) did not study this protein further for iron-dependent regulation. Additionally, CC3336,
120 a protein that my study identified as iron regulated, was identified by Landt (13) as being carbon regulated. CC3336, and potentially CC3161, appear to be regulated by both iron and carbon.
In a study by Landt et al. (13), the transcript for CC3336 was stabilized during carbon starvation by a sRNA called CrfA. CrfA has been seen to affect the expression of several genes in this way (13). It interacts with a stem-loop in the 5’
UTR, stabilizing the RNA as shown in Figure 4.2 (13). This, combined with my data, indicate that CC3336 is under the regulation of two different nutrients: iron and carbon.
I identified CC3336 as being up regulated under iron starvation conditions at the mRNA level but could not locate any sequences in the promoter area that resembled a Fur box or an IRO (iron response operator). There is a chance that the iron regulation of this gene could be taking place post-transcriptionally. For one thing, there is most likely a structure in the 5’ UTR that enables this mRNA to interact with a sRNA in a positive, stabilizing fashion (13). Fur has been seen to have the ability to induce expression of genes in response to adequate iron by repressing small regulatory RNAs that would normally repress these genes (2, 5,
14, 19, 24). Hence, it is known that some iron regulation has been seen to take place post-transcripionally. It is not possible at this point to tell if the iron regulation of CC3336 is occurring transcriptionally or post-transcriptionally, but the basic scheme of CC3336 regulation as I now know it is illustrated in Figure
4.2. If the iron-dependent regulation is taking place post-transcriptionally, this
121
Figure 4.2: Two possible scenarios for the regulation of CC3336. In both cases,
CC3336 mRNA is stabilized by carbon starvation (13). When the cell is carbon starved, the sRNA
CrfA stabilizes the mRNA most likely by binding to the 5’ UTR. This results in stabilization so that the mRNA can be translated into an outer membrane receptor. A. Iron starvation results in a post- transcriptional response mediated by an unknown regulator, destabilizing the mRNA. B. Iron starvation causes an iron-dependent regulator not to bind to the promoter. This allows CC3336 to be transcribed.
122 would be an interesting feature, because this would indicate that this mRNA is
stabilized in one condition and destabilized another. Because CC3336 is not
listed as a gene regulated by Fur in the study by da Silva Neto et al. (3), it is
probably either being iron regulated by another transcription factor such as
CC2625 or by a sRNA. If CC2625 is a novel regulator, it would likely have a
different promoter binding site than the previously studied regulators and I would not have known what consensus sequence to look for in the promoter of CC3336 when it was being examined. Interestingly, no other genes besides CC3336 were identified by my experiments or those done by da Silva Neto et al. (3) as being iron regulated and also identified by Landt et al. (13) as being carbon regulated.
CC3161 was found to be carbon regulated, and in the study done by da Silva
Neto et al. (3), it was predicted to have a weak Fur binding site, but they did not confirm any iron-dependent regulation. Also, neither of the TonB-dependent receptors with known fuction (i.e. carbohydrate uptake) were mentioned by Landt et al. (13) as being carbon regulated by CrfA (7, 13, 17, 18) . This overlap in carbon and iron regulation has not been reported in any other known genes in C. crescentus.
It seems that CC3336 is controlled by different forms of starvation.
Perhaps this could be some complicated mechanism of changing cellular metabolism based on the nutrients available. Several known TonB-dependent iron receptors in E. coli are regulated by both iron and carbon at the transcriptional level by Fur and Crp (15, 28). It is not known why these genes are
123 regulated in this fashion (15, 28). It is possible that CC3336 is an iron receptor which is activated during carbon starvation in order to bring in more iron. That seems like a strange action for a cell to take. However, it is possible that a metabolic “decision” is being made here. Of course, there are many metabolic processes where carbon is needed. Perhaps, under carbon starvation conditions, there is an advantage for the cell to bring in more iron, and it is helpful to the cell
in overcoming a lack of carbon. However, at this time, the reasons for this type of
regulation are not yet apparent.
Dual regulation of receptors is a very exciting direction for future research.
A good place to start would be to grow the CC3336 mutant under more iron and
carbon starvation conditions in order to obtain a growth phenotype and to examine the OMP profile of this strain in different growth conditions. In addition, it may also become important to take a closer look at CC3161 which was identified both as being carbon regulated and having a weak Fur binding site (3, 13). It
would first be important to examine this protein to see if iron regulation is
apparent. If it is, CC3161 is also a candidate for the study of this dual regulation
phenomenon.
Overall, through my receptors study, nine putative iron regulated TonB- dependent receptors were identified. Further work will need to be done in order to confirm these identifications, to clarify the level of iron regulation for many of these, and also to match up receptors with the type of ferric-siderophore they bring into the cell. A good place to begin future studies in this area would be
124 through the construction of deletions in each of these ORFs, and also in combinations of ORFs, in order to cut down on the likelihood of gene
redundancy. New mutants could be grown on various siderophores in order to
determine the cognate siderophore for each receptor.
Conclusion
Through the study of metal-dependent regulators and iron regulated
TonB-dependent receptors, a better understanding of the metal-dependent
regulation system in C. crescentus has been attained. More work is needed in
this area in order to compose the entire picture. I have identified a putative novel
regulator of the Rrf2 family of proteins and have demonstrated that it is metal-
dependent. I have also identified a few iron regulated TonB-dependent receptors,
including one with a dual regulation that has not been seen before. This work has
established some of the first puzzle pieces in the overall metal-dependent uptake
and regulation system in this microorganism. Understanding this particular
system in C. crescentus could lead to a better comprehension of similar systems
in other aquatic bacteria. Since aquatic bacteria have been little studied in this
area, this research could provide insight into the inner-workings of a novel and
complex system of regulation.
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