INSIGHTS INTO MICROBIAL METABOLISM
by
Mary Catherine Burgess
A thesis submitted in partial fulfillment
of the requirements for the degree
of
Master of Science
in
Biochemistry
MONTANA STATE UNIVERSITY
Bozeman, Montana
April 2011
©COPYRIGHT
by
Mary Catherine Burgess
2012
All Rights Reserved
ii
APPROVAL
of a thesis submitted by
Mary Catherine Burgess
This thesis has been read by each member of the thesis committee and has been
found to be satisfactory regarding content, English usage, format, citation, bibliographic
style, and consistency and is ready for submission to The Graduate School.
John W. Peters
Approved for the Department of Chemistry and Biochemistry
Bern Kohler
Approved for The Graduate School
Dr. Carl A. Fox
iii
STATEMENT OF PERMISSION TO USE
In presenting this thesis in partial fulfillment of the requirements for a master’s
degree at Montana State University, I agree that the Library shall make it available to
borrowers under rules of the Library.
If I have indicated my intention to copyright this thesis by including a copyright
notice page, copying is allowable only for scholarly purposes, consistent with “fair use”
as prescribed in the U.S. Copyright Law. Requests for permission for extended quotation
from or reproduction of this thesis in whole or in parts may be granted only by the
copyright holder.
Mary Catherine Burgess
April 2012
iv
ACKNOWLEDGEMENTS
I would like to first thank my advisor, John Peters, for his support in this endeavor and his
willingness to work with me on several different projects. I would also like to thank my friends
and family who have been extremely supportive of me during this process, especially my mom
and Mike who have always supported my best interests and have always had time to talk and
help me through the difficult and frustrating times. I would also like to thank all of the members
of the Rae and Sourdough Fire Departments who have, for three years, given me a reprieve
from academia whenever I needed it.
I appreciate all of the members of the Peters’ lab: Eric, Trinity, Oleg, Dan, Jesse, Kevin,
Neelambari, Steve, Alta, Wes and Matt who have provided help and advice when I have needed
it and made working in the lab enjoyable. I would particularly like to thank Eric for his help with
my environmental projects and being patient while I learned to work with environmental
samples and providing valuable insights into the projects. I am extremely grateful to Trinity who
has spent countless hours of her time showing me the ropes of the lab from day one and helping
me with troubleshooting my projects and reviewing my papers. I would also like to thank
countless other faculty and graduate students in the Chemistry and Biochemistry Department
for being willing to take time out of their days to lend me their advice and expertise while
working through difficult projects.
v
TABLE OF CONTENTS
1. DEFINING THE FUNCTIONAL ROLE OF NITROGENASE
HOMOLOGS IN ROSEIFLEXUS CASTENHOLZII USING
STRUCTURAL STUDIES ................................................................................................................. 1
Introduction................................................................................................................................. 1
Nitrogenase Reaction and Structure .................................................................................. 1
Nitrogenase Iron Protein ....................................................................................... 2
Nitrongease MoFe Protein .................................................................................... 4
Photosynthesis ................................................................................................................... 9
Roseiflexus castenholzii as a model organism .................................................................. 12
Materials and Methods ............................................................................................................. 15
Growth without fixed N .................................................................................................... 15
Development of an expression and purification
protocol for nitrogenase homologs.................................................................................. 16
Protein Expression in E. coli and Purification ..................................................... 16
Roseiflexus castenholzii Nitrogenase NifH
homolog Purification .......................................................................................... 17
Roseiflexus castenholzii Nitrogenase
MoFe-homolog Protein Purification ................................................................... 18
Roseiflexus castenholzii nifB gene
Product Purification ............................................................................................ 19
Results and Discussion .............................................................................................................. 19
Growth of Roseiflexus castenholzii without fixed N ......................................................... 19
Expression and Purification of Roseiflexus castenholzii
Iron Protein Homolog ....................................................................................................... 20
Expression and Purification of Roseiflexus castenholzii
MoFe Protein Homolog .................................................................................................... 21
Expression and Purification of Roseiflexus castenholzii
Nif B Protein Homolog...................................................................................................... 24
References ................................................................................................................................. 25
2. EXPRESSION AND PURIFICATION OF THREE CYANOBACTERIAL
TRICARBOXYLIC ACID CYCLE PROTEINS ...................................................................................... 30
Introduction............................................................................................................................... 30
The Tricarboxylic Acid Cycle Overview ............................................................................. 30
The Tricarboxylic Acid Cycle in Eukaryotes ...................................................................... 32
The Tricarboxylic Acid Cycle in Prokaryotes ..................................................................... 33
Materials and Methods ............................................................................................................. 40
Growth of select Synechococcus sp.
PCC7002 TCA enzymes ..................................................................................................... 40
Purification of select Synechococcus sp.
PCC7002 TCA enzymes ..................................................................................................... 41
Results and Discussion .............................................................................................................. 43
vi
TABLE OF CONTENTS - CONTINUED
Fumarase .......................................................................................................................... 43
Succinic Semialdehyde Dehydrogenase ........................................................................... 45
2-Oxoglutarate Decarboxylase ......................................................................................... 48
References ................................................................................................................................. 50
3. ECOLOGICAL SIGNIFICANCE OF A SULFATE RESPIRING
MICROORGANISM FROM AN ACID-SULFATE-CHLORIDE
GEOTHERMAL FEATURE IN
YELLOWSTONE NATIONAL PARK ................................................................................................ 54
Introduction................................................................................................................................ 54
The Mercury Cycle ............................................................................................................ 54
Methyl Mercury in Yellowstone National Park ................................................................. 60
Materials and Methods .............................................................................................................. 66
Sample Collection and Enrichment Conditions ................................................................ 66
Epifluorescence Microscopy............................................................................................. 67
PCR Analysis...................................................................................................................... 68
16S rDNA Gene Analysis ................................................................................................... 69
Results and Discussion ............................................................................................................... 69
Dragon Spring Site Description......................................................................................... 69
Dragon Spring Isolate Enrichment and Isolation .............................................................. 69
Morphology of Dragon Spring Culture Sample ................................................................ 72
References .................................................................................................................................. 75
4. CONCLUSIONS ............................................................................................................................ 82
REFERENCES............................................................................................................................... 85
vii
LIST OF FIGURES
Figure
Page
1.1 Structure of the nitrogenase iron protein .................................................................... 3
1.2 Structure of the nitrogenase MoFe protein.................................................................. 5
1.3 P cluster structures ....................................................................................................... 6
1.4 FeMo-cofactor structure ............................................................................................... 7
1.5 SDS-PAGE of R. castenholzii iron
protein purification..................................................................................................... 21
1.6 Initial NifD and NifK purification ................................................................................. 22
1.7 NifD and NifK purification gels .................................................................................... 23
2.1 The Tricarboxylic Acid Cycle ........................................................................................ 31
2.2 Interactions of metabolic pathways ........................................................................... 32
2.3 The glyoxylate cycle .................................................................................................... 34
2.4 Conversion of α-ketoglutarate to succinate ............................................................... 37
2.5 Proposed alternative TCA cycle in
Synechococcus sp. PCC 7002 ....................................................................................... 39
2.6 SDS-PAGE of fumarase purification ............................................................................ 43
2.7 Western blot of fumarase Ni2+-NTA elutions .............................................................. 43
2.8 SEC peak of fumarase ................................................................................................. 44
2.9 SDS-PAGE gel of SSADH purification ........................................................................... 46
2.10 Western blot of SSADH Ni2+-NTA elutions ................................................................ 46
2.11 SEC peak of SSADH .................................................................................................... 47
2.12 SDS-PAGE of 2-OGDC purification............................................................................. 48
2. 13 Western blot of 2-OGDC Ni2+-NTA elutions ............................................................. 49
viii
LIST OF FIGURES – CONTINUED
Figure
Page
3.1 Mercury metabolism by microorganisms ................................................................... 58
3.2 Methyl mercury complexed with cysteine ................................................................. 59
3.3 Abiotic and biotic reactions of the sulfur cycle........................................................... 61
3.4 Dragon spring sampling site........................................................................................ 64
3.5 Map of Yellowstone National Park ............................................................................. 65
3.6 PCR amplicons of bacterial 16S genes ........................................................................ 70
3.7 Epifluorescent image of Dragon
Spring enrichment culture .......................................................................................... 73
ix
ABSTRACT
Nitrogen fixation (catalyzed by the enzyme nitrogenase), cellular respiration (completed
through the Tricarboxylic Acid (TCA) cycle) and mercury detoxification (through mercury
methylation) are three metabolic processes used by a wide variety of microorganisms, but that
also have far reaching impacts on nutrient cycling in the environment. Roseiflexus castenholzii
has been found to have a unique nitrogenase gene cluster encoding several nitrogenase
homologs, including the structural proteins NifH and NifDK and the radical SAM protein, NifB,
necessary for cofactor biosynthesis. However, the genome of R. castenholzii lacks the suite of
nitrogenase accessory proteins necessary for nitrogen fixation. To investigate the metabolic role
of these nitrogenase homologs, expression and purification protocols were developed that aid
in the biochemical characterization of these proteins. Synechococcus sp. PCC 7002 encodes
three novel TCA proteins, contrary to previous studies that indicated these phototrophs have
incomplete TCA cycles. Expression, purification and preliminary crystallization trials were
completed on the three novel TCA proteins in order to gain insight into the structure of the
proteins which will elucidate the mechanism of each novel enzyme and provide evidence into
the novel TCA cycle utilized by these cyanobacteria. The third project presented examines the
role of microorganisms in metabolizing mercury, producing methylmercury and providing an
entry point for methylmercury into the food chain in Yellowstone National Park (YNP). In this
project, environmental samples were enriched for a sulfate reducing organism and a culture
containing three sulfate reducing bacteria (SRB) has been established. The SRB that are present
and active in the enrichment samples are known to reduce sulfate and may be responsible for
the presence of methyl mercury in algal mats that bioaccumulates through the food chain in
YNP. The enrichment of SRB in this culture will enable the identification and characterization of
the organisms that are capable of methylating mercury in hydrothermal systems. Collectively,
the results presented herein increase the knowledge base of three metabolic processes used by
microorganisms: nitrogen fixation, cellular respiration through the TCA cycle and mercury
detoxification; these results will contribute to a broader understanding of how these processes
have evolved and their impacts on the environment.
1
CHAPTER 1
DEFINING THE FUNCTIONAL ROLE OF NITROGENASE HOMOLOGS IN ROSEIFLEXUS CASTENHOLZII
USING STRUCTURAL STUDIES
Introduction
Nitrogenase Reaction and Structure
Nitrogenase is a two-component metalloenzyme that catalyzes the ATP-dependent
conversion of dinitrogen to ammonia:
N2 + 16MgATP + 8e- + 8H+→2NH3 + H2 + 16 MgADP + 16Pi
Without this enzyme and several abiotic sources of fixed nitrogen, the most abundant form of
nitrogen in the atmosphere, dinitrogen, would be bio-unavailable. As the only biotic source of
fixed nitrogen, it is estimated that nitrogenase produces more than 2 x 1013g of fixed nitrogen
per year [3]. The most predominant abiotic sources of fixed nitrogen are lightening strikes which
produce approximately 1012g/year; and the Haber-Bosch process which produces ammonia
industrially. Unlike the reaction performed by nitrogenase at physiological temperatures and
pressures, the Haber-Bosch process requires a significant amount of energy, with catalysis
occurring at 400-5000C and 250 atm, while yielding only 10-20% ammonia [4, 5]. In contrast to
the energetically expensive Haber-Bosch process, biological nitrogen fixation by nitrogenase is
gaining recognition as a more sustainable option. The nitrogenase enzyme is found only in
prokaryotes that inhabit a wide variety of environments, both terrestrial and marine [6, 7].
Nitrogenases are found in both aerobic and anaerobic organisms, even though the characteristic
iron-sulfur clusters of the enzyme are oxygen sensitive. The ability of nitrogenases to exist in
oxygenic environments is due to several adaptations including spatial separation of the iron-
2
sulfur clusters from oxygen (by separating nitrogenases from oxygenic activites) and temporal
separation of processes requiring oxygen from the iron sulfur clusters of the nitrogenase (for
example, fixing nitrogen at night and completing photosynthesis during the day) [8, 9].
There are three different kinds of characterized nitrogenase enzymes named based on
the metal composition of their active site: molybdenum-dependent nitrogenase (nif, the most
abundant, and the focus of this project), vanadium-dependent nitrogenase (vnf) and iron only
nitrogenase (anf). All known, functional nitrogenases exist as two component enzymes: the
dinitrogenase reductase (Fe protein) and the dinitrogenase (“MoFe protein” in the
molybdenum-dependent nitrogenase). There are also uncharacterized nitrogenases (with
unknown active site compositions) and nitrogenase homologs, which are characterized as such
based on sequence similarity.
Nitrogenase Iron Protein
The Fe protein, the nifH gene product, is a homodimer bridged by a [4Fe-4S] cluster
coordinated by four cysteine residues (two from each monomer) (Figure 1.1) [10]. The ironsulfur cluster is located symmetrically between the dimers of the iron protein, a bridging
structure that is unique in biology. The L protein of the protochlorophyllide reductase (BchL)
involved in photosynthesis also has a symmetric bridging iron-sulfur cluster and is evolutionarily
related to the nitrogenase iron protein [11]. Sequence and structural homology between the
two proteins led to the hypothesis that they evolved from a common ancestor.
3
Figure 1.1. Nitrogenase Fe protein. NifH monomers are red and green,
and the bridging Fe-S cluster is indicated with an arrow PDB ID 1CP2.
In the nitrogenase Fe protein hydrogen bonds stabilize the iron-sulfur cluster within
the protein [12]. The iron protein’s main function is to transfer electrons from reduced
ferredoxin or flavodoxin to the MoFe protein. Complete reduction of the substrate N2 requires
six electrons which are transferred individually from the Fe protein to the MoFe protein. This
electron transfer is indirectly coupled to the hydrolysis of MgATP, which makes the iron protein
unique, since most electron transfer proteins do not require ATP [10, 13]. The MgATP binds
approximately 20Å from the iron-sulfur cluster and is thought to be required for a
conformational change in the Fe protein, similar to changes seen in nucleotide-dependent signal
transduction proteins [14]. Through structural comparisons of the MgADP bound and unbound
structures of the protein as well as mutational studies involving residues at the Fe-MoFe protein
interface, it was found that there are conformational changes at the Fe protein-MoFe protein
4
interface associated with MgATP hydrolysis [15-19]. The iron protein also functions in the
biosynthesis of the FeMo cofactor, a complex metal cofactor where substrate reduction occurs
in the MoFe protein.
Due to the sensitivity of the iron protein’s iron-sulfur cluster to oxygen, all studies of
the protein must be completed anaerobically. Because of the difficulty of working with the
protein, and the fact that several organisms (including A. vinelandii) have Fe proteins that are
not heterologously expressed well in E. coli, the exact mechanism of catalysis of the iron protein
has been difficult to study. However, several studies of the A. vinelandii Fe protein have been
completed which illuminate unique characteristics of the protein. The iron-sulfur cluster of the
Fe protein exists in three redox states: the most common [4Fe-4S]+2 (oxidized state), the [4Fe4S]+1 (reduced state), and the least common [4Fe-4S]0 state (all ferrous form). Strop et al, solved
the structure of the all ferrous [4Fe-4S]0 form of the nitrogenase iron protein, and found that
the iron-sulfur cluster has the largest surface area exposure of any published [4Fe-4S] clustercontaining protein [12]. The surface area exposure ranges from 13.5-18.2Å2 in the all-ferrous
form of the Fe protein (PDB ids: Av2, 1G5P, 1CP2). They propose that the solvent exposure of
the cluster facilitates reduction of the cluster and may relate directly to the ability of the protein
to exist in three oxidation states [12].
Nitrogenase MoFe Protein
The nitrogenase MoFe protein, composed of nifD and nifK gene products, forms an
α2β2 heterotetramer with two different complex iron-sulfur clusters (and two of each type)
(Figure 1.2). The nifD and nifK genes share sequence similarity due to their emergence from an
ancient gene duplication event [20, 21]. Each α subunit of the MoFe protein binds to a β subunit
and forms a distinct dimer; therefore in the active heterotetramer the MoFe protein has a two-
5
fold symmetry between equivalent dimers. The αβ dimers of the MoFe protein interact solely
through interactions between the β subunits. In each dimer there are two different types of
iron-sulfur clusters: a P cluster and a FeMo cofactor [22]. Thus, in the MoFe protein there are a
total of four complex iron-sulfur clusters. Electrons are transferred from the Fe protein to the P
cluster of the MoFe protein and finally to the FeMo-cofactor of the MoFe protein which is the
active site where the substrate N2 is bound and reduced.
Each P cluster is composed of two [4Fe-4S]-like subclusters bridged by a
hexacoordinate sulfur and bound to the protein by six cysteine ligands [22-26]. The complex and
Figure 1.2. Nitrogenase MoFe protein, with arrows
indicating the sites of the metal clusters in one dimer.
Red (β subunit); Yellow (α subunit) PDB ID 3U7O.
6
unique structure of the P cluster made solving its structure and identifying the form of the
cluster present in the active MoFe protein difficult (Figure 1.3) [23, 26].
Figure 1.3. P cluster structure (Fe atoms-red; S atomsyellow).
The P cluster is assembled in the MoFe protein, unlike the FeMo-cofactor which is
assembled on scaffold proteins and then ligated into the MoFe protein [27]. Because the P
cluster is assembled in the MoFe protein, it was difficult to differentiate the form of the P cluster
present in the active protein. Studies of the cluster have shown evidence of two pairs of [4Fe4S]-like clusters and two [8Fe-7S] clusters. Discerning which type of cluster is present in the
active protein, versus which cluster is a precursor was difficult. Additional studies found that the
cluster composition is redox dependent [26], and the [4Fe-4S]-like clusters are precursors to the
[8Fe-7S] clusters present in the mature protein [27]. The P cluster is located at the αβ dimer
interface and between the [4Fe-4S] cluster of the Fe protein and the FeMo-cofactor where it
functions to transfer electrons from the iron protein to the FeMo-cofactor. The oxidationreduction potential of the P cluster is sensitive to pH which indicates that the P cluster may
couple electron transfer with proton uptake and transfer [26]. Maturation of the MoFe protein
7
progresses in stages. In the first stage the P cluster is formed, then there is a lag time after
which the second P cluster is formed. Both P clusters must be present before the FeMocofactor can be inserted [27].
The FeMo-cofactor (FeMo-co) is the second type of iron sulfur cluster found in the
nitrogenase MoFe protein. Located entirely in the α subunit and composed of a [7Fe-9S-Mo-Chomocitrate] cluster, the cofactor serves to accept electrons from the P cluster and bind
dinitrogen to reduce it to ammonia (Figure 1.4) [22, 28]. The carbon is a central atom in the
cluster that has been unidentified for many years. It was predicted to be a carbon, oxygen or
nitrogen, recently predicted to be a carbon [29] and confirmed to be carbon through x ray
emission spectroscopy and x-ray crystallography combined with radiolabelling [30-32]. The
FeMo-co is anchored to the protein through covalent coordination through a cysteine (with an
Fe atom) and a histidine (with the Mo atom). There is also a homocitrate molecule linked to
FeMo-co which is coordinated to the Mo atom and linked to the protein through hydrogen
bonds and water-bridged hydrogen bonds [25].
C
Mo
Homocitrate
Figure 1.4. FeMo cofactor (Sulfur-yellow; iron-orange;
carbon-green; oxygen-red; molybdenum-blue).
8
The FeMo cofactor in the MoFe protein is clearly very comples and requires a large
suite of gene products to be present to assist in synthesis of the cofactor and maturation of the
MoFe protein (reviewed in [33, 34]). Biosynthesis of the FeMo cofactor requires NifU, NifS, NifH,
NifZ, and NafY. NifU and NifS are a molecular scaffold and a cysteine desulfurase, respectively,
which provide Fe-S clusters for nitrogenase-specific proteins [35]. NifZ has been proposed to be
a chaperone involved in formation of the second P cluster [36]. NafY is responsible for delivering
FeMo-cofactor to the apo-MoFe protein and stabilizing the apo-MoFe protein so that it can bind
the cofactor [37]. Previous studies of nif mutant strains, each lacking the gene for one or more
accessory protein, resulted in partially or completely inactive MoFe proteins [38]. In addition to
the proteins needed for MoFe protein maturation, there are several accessory proteins needed
for FeMo-cofactor biosynthesis including: NifU, NifB, NifQ, NifEN, NifX, NafY, NifS, and NifV [33].
In this project only the role of NifB and NifEN will be addressed. It should be noted, however,
that several of the other accessory proteins are necessary for FeMo-cofactor biosynthesis in
vivo, and several others are only required under certain metabolic conditions [38].
NifB uses iron-sulfur clusters provided by NifU and NifS to form NifB-co in a Sadenosyl methionine (SAM) radical reaction [39]. NifU and NifS are involved in iron and sulfur
sequestration and iron-sulfur cluster assembly, and also are homologs to proteins found in the
ISC and SUF iron-sulfur cluster assembly systems. In deletion strains lacking NifB, the MoFe
protein lacks a FeMo-cofactor and electrons cannot be transferred to reduce N2, rendering the
nitrogenase inactive [37]. NifEN is a scaffold protein that aids in FeMo-cofactor assembly and
may be the result of a nifDK gene duplication event, suggested by sequence similarities between
the proteins [40]. An in vitro system containing the NifEN, NifB, and Fe protein as well as
homocitrate, MgATP and S-adenosyl methionine is the most minimal system to date that is able
9
to assemble a complete FeMo-cofactor and deliver it to the MoFe protein [38]. This study, and
the fact that not all diazotrophs require the full complement of accessory proteins suggests that
proteins from other iron sulfur cluster assembly systems such as the iron-sulfur cluster (ISC) or
the sulfur assimilation (SUF) systems may be able to substitute in some prokaryotes (reviewed in
[41]). The ISC and SUF systems are general pathways for assembly of a variety of proteins
containing iron-sulfur clusters. Also, the gene products of the ISC and SUF systems function
similarly to some of the gene products involved in nitrogenase maturation. For example, the
complex, SufBCD functions in an ATP dependent manner to assist the complex SufSE (with
cysteine desulfurase activity) in building iron-sulfur clusters and delivering them to the scaffold
protein, SufA [41]. However, it should be noted that results from the in vitro study of a minimal
system that can synthesize an active FeMo-cofactor may not indicate what is needed for an in
vivo system; nifU or nifS, and possibly other gene products may be critical components of in
vivo FeMo-cofactor biosynthesis.
The nitrogenase MoFe protein, like the Fe protein, is evolutionarily related to a
protein necessary for photosynthesis, the NB-protein which is involved in chlorophyll
biosynthesis [42]. The NB-protein, which is discussed in further detail below, is a BchN-BchB
heterotetramer which reduces porphyrin using iron-sulfur clusters. Although the clusters are
located in spatially similar places to those found in the MoFe protein, the NB-protein only
contains [4Fe-4S] clusters.
Photosynthesis
Photosynthesis is a chemical process in which water and carbon dioxide react to form
organic compounds and oxygen. The general reaction is always the same:
carbon dioxide + electron donor + light energy → carbohydrate + oxidized electron donor
10
However, different organisms may use different electron donors or acceptors. The
photosynthesis reaction requires several different components which include photosynthetic
pigments, reaction centers, electron transport chains, and antenna systems. The two most
common kinds of light reactive pigments are called bacteriochlorophyll (Bchl, found in
prokaryotic organisms) and chlorophyll (Chl, found in plants, algae and cyanobacteria), both are
synthesized from a tetrapyrrole ring system. Reaction centers (which are made up of several
proteins, pigments and cofactors and are the location where light energy reacts to excite
electrons) are divided into two types: type I and type II which are distinguished by the electron
acceptor cofactors in their active site [43]. In type I reaction centers the electron acceptor
cofactors are iron sulfur clusters, whereas in type II reaction centers the electron acceptor
cofactors are pheophytin/quinine complexes. Electron transport chains are made of cytochrome
complexes located within the reaction centers. Several cytochrome complexes are known to
exist, the most common being bc1 and b6f, and in filamentous anoxygenic phototrophs, complex
III. The final component needed for photosynthesis is an antenna system. Antenna systems
function to gather light and transfer the light energy to the reaction center. Many diverse
antenna systems exist with different pigments and structures that are optimized for an organism
and its environment. Of key interest to this project is the biosynthetic machinery that is
evolutionarily related to nitrogenase proteins involved in synthesizing the light reactive
pigments of photosynthesis, Bchl and Chl. Chlorophyll biosynthesis, like nitrogenase
biosynthesis, is a complex process with many steps and accessory proteins. During Chl/Bchl
synthesis, a molecule called protochlorophyllide (Pchlide) is reduced at the C-17=C-18 double
bond to make chlorophyllide (Chlide). Chlide is then made directly into chlorophyll a or, in the
11
case of bacteriochlorophyll biosynthesis Chlide is further reduced to form bacteriochlorophyllide
from which bacteriochlorophyll is produced.
One widely accepted theory of chlorophyll evolution that is based on the Granick
hypothesis states that Bchl evolved after Chl because Chl synthesis requires fewer enzymatic
reductions [11]. As mentioned previously, the bacteriochlorophyll and chlorophyll biosynthetic
machinery shares sequence similarity with nitrogenase proteins. Light-independent (dark
operative) protochlorophyllide oxidoreductase (DPOR) is one of two enzymes responsible for
reducing the C-17=C-18 double bond of ring D of protochlorophyllide chlorophyllide. Plants,
algae and cyanobacteria have an alternative enzyme that reduces the double bond: lightdependent Pchlide oxidoreductase (LPOR) which shares no sequence similarity with DPOR.
DPOR is comprised of three subunits, BchN, BchB and BchL in bacteriochlorophyll-synthesizing
systems (or ChlN, ChlB and ChlL in chlorophyll-synthesizing systems). The overall structure of the
DPOR enzyme is similar to the nitrogenase enzyme in that it is composed of BchL (or ChlL), a
homodimer which binds to a BchNB (or ChlNB) heterotetramer that resembles the binding of
the NifH homodimer to the NifDK heterotetramer. The DPOR proteins N, B and L are
evolutionarily related to NifD, NifK and NifH, respectively.
There is also a second protein, chlorophyllide oxidoreductase (COR), composed of
subunits BchY, BchZ and BchX which is unique to bacteriochlorophyll systems because it reduces
the C-7=C-8 double bond of chlorophyllide a to form bacteriochlorophyllide a (the precursor of
bacteriochlorophyll a) [2]. COR has an overall structure very similar to DPOR: BchX forms a
homodimer that binds to a heterotetramer of BchYZ (Figure 1.6). The structures and functions of
the COR component proteins are similar to their nitrogenase homologs in that BchX is an ATP
dependent reductase with an iron-sulfur cluster and the BchYZ protein contains the active site
12
and two iron-sulfur clusters per dimer [44]. The subunits of COR also have sequence similarity to
NifD, NifK and NifH. The similar structures and chemistries of the (bacterio) chlorophyll
biosynthesis enzymes DPOR and COR to nitrogenase provides evidence for historic gene
duplication events or divergence from a common ancestor.
The BchL protein has 33% amino acid sequence identity with NifH [11]. Like NifH, BchL
is a homodimer with a bridging [4Fe-4S] cluster coordinated by four cysteine residues at the
dimer interface and one MgATP binding site per monomer [45]. Hydrolysis of the MgATP results
in structural changes in the BchL protein (as it does in NifH) that occur along the BchL/NBprotein interface. The NB-protein of DPOR contains two Pchlide binding sites and two NBclusters, one of each in each catalytic BchN-BchB subunit. The NB cluster is a [4Fe-4S] cluster
coordinated to the protein through one aspartate and three cysteine residues [42]. Mutational
studies have shown that the aspartate ligand is not critical for assembly of the NB-cluster, but
does play a role in enzymatic activity [42]. Unlike the MoFe protein, the NB-protein does not
have two different kinds of clusters. Instead, electrons are transferred from the NB cluster
directly to the substrate Pchlide for reduction. The reduction of Pchlide is complex and requires
the trans-specific reduction of the C-17=C-18 double bond which is possible because of the
distorted configuration of the C-17-propionate of Pchlide [42].
Roseiflexus castenholzii as a Model Organism
Roseiflexus castenholzii is a filamentous anoxygenic phototroph (FAP) isolated from a
Japanese hot spring. R. castenholzii grows optimally at 50oC, pH 7.5 and chemoautotrophically
under dark aerobic conditions or photoheterotrophically under light anaerobic conditions. R.
castenholzii contains bacteriochlorophyll a, the genome encodes nitrogenase homologs nifH,
nifB, nifD, and nifK, but lacks chlorosomes and bacteriochlorophyll c and any additional
13
nitrogenase accessory proteins. The role of the limited nitrogenase gene cluster in R.
castenholzii is not known; hypotheses have been made that the nitrogenase homologs may be
involved in photosynthesis or nitrogen fixation. As a member of the family Chloroflexaceae, R.
castenholzii is important in the origin and evolution of photosynthesis because members of the
phylum Chloroflexi are the earliest branching phylum of bacteria which contains phototrophs
[46, 47]. Recent phylogenetic analyses of Nif, Vnf, Anf, uncharacterized nitrogenases and
nitrogenase homologs place the R. castenholzii nitrogenase homologs in a lineage which
branches after Nif, the alternative nitrogenases and the uncharacterized nitrogenases [48].
These studies suggest that the nitrogenase homologs from the filamentous anoxygenic
phototrophs (including R. castenholzii) are the most recently evolved lineage of putative
nitrogenases.
Some preliminary studies have characterized the light harvesting reaction complex of
R. castenholzii and unpublished results from the Bryant lab suggest that the nitrogenase
homologs are not being upregulated during photosynthesis, suggesting the homologs may not
be involved in photosynthesis; but to date no studies on the nitrogenase homologs have been
performed [49]. The nitrogenase homologs present in R. castenholzii have not been
characterized, and thus are not known to possess any nitrogenase activity; although
phylogenetic analyses place them with known nitrogenases [48]. An active nitrogenase with only
the nifH, nifB, nifD and nifK gene products is unprecedented; the role of the nitrogenase
homologs in R. castenholzii is unknown. Nif-like proteins homologous to NifH and NifD but with
no nitrogenase activity have been identified in all methanogens and are hypothesized to be
necessary for growth of the methanogens. Therefore, it is possible that the R. castenholzii
14
nitrogenase homologs have no nitrogenase activity, but are necessary for the growth of the
organism [50].
Although phylogenetic analyses place the R. castenholzii nitrogenase homologs in a
lineage with the most recently evolved putative nitrogenases, whether the gene products could
function as a nitrogenase without having the genes necessary for cofactor biosynthesis and
insertion remains a question. Other possible roles for these proteins could be in
bacteriochlorophyll biosynthesis, because as mentioned previously, bacteriochlorophyll
biosynthesis proteins share homology with nitrogenase proteins. However, unpublished
preliminary results from Dr. Donald Bryant do not suggest a bacteriochlorophyll biosynthesis
function for the R. castenholzii nitrogenase homologs; it was observed that under conditions
promoting bacteriochlorophyll biosynthesis the nitrogenase homologs were not up-regulated
(as would be expected if they had a function in bacteriochlorophyll biosynthesis).
The goal of this project is to investigate the expression of the R. castenholzii
nitrogenase homologs when grown heterologously in E. coli in order to determine if
heterologous expression is a tractable system for characterization of the homologs. Extensive
research completed on nitrogenase enzymes from model organisms such as A. vinelandii will be
used as a guide to complete this project. Work was completed recently by Curatti, et. al. to
establish a system using the minimal number of nitrogenase accessory proteins and reactants
such as MgATP and S-adenosyl methionine necessary to achieve reduction of dinitrogen by the
nitrogenase enzyme [38]. They characterized a system with only NifEN, NifB, and NifH as well as
homocitrate, MgATP and S-adenosyl methionine which could assemble a complete FeMocofactor and deliver it to the MoFe protein in vitro. These results suggest that if such a limited
system could function in vitro to assemble a FeMo-cofactor, then perhaps in organisms lacking a
15
complete compliment of nitrogenase accessory proteins alternative assembly proteins can be
used; it is possible that proteins from other iron sulfur cluster assembly systems such as the ISC
or SUF systems are substituting for traditional nitrogenase accessory proteins. The genome of R.
castenholzii does contain the SUF system genes. In addition to the results of these studies, it has
been found that some methanogens are able to fix nitrogen without having all of the
nitrogenase accessory genes. Methanogens have been shown to have only the nifHDKENBV
genes [48]. Because of these results it is possible that the nitrogenase homologs in R.
castenholzii could fix nitrogen; the role of the nitrogenase homologs as well as a system for
heterologous expression of the homologs is investigated in this study.
Materials and Methods
Growth Without Fixed Nitrogen
R. castenholzii was first isolated and characterized by Hanada, et. al., in 2002 [51]. Their
results show that R. castenholzii is maintained on PE medium with yeast extract, under both
aerobic dark and anaerobic light conditions. PE medium contains (per liter): 0.5 g sodium
glutamate, 0.5 g sodium succinate, 0.5 g sodium acetate, 0.5 g yeast extract, 0.5 g Casamino
acids, 0.5 g Na2S2O3·5H2O, 0.38 g KH2P04, 0.39 g K2HP04, 0.5 g (NH4)2S04, 1 ml vitamin mixture,
and 5 ml basal salt solution (for further information on PE media see [52]). During isolation,
decreased growth with citrate, lactate, glucose, and Casamino acids was observed [51].
For evaluation of the potential for R. castenholzii to grow diazotrophically, PE medium
was altered to contain carbon sources (0.25%w/v) that do not contain fixed N, such as: acetate,
citrate, glucose, glycyl-glycine, and pyruvate. Media lacking fixed N was inoculated with cultures
grown on fixed N in late log phase (OD600 = 0.6). After 64 hours of growth, cultures were
16
transferred into the same media lacking fixed N in order to remove any fixed N that may have
been present from the original inoculums. Growth in minimal media lacking fixed N with varying
carbon sources was monitored by OD600.
Development of an Expression and
Purification Protocol for Nitrogenase Homologs
Protein Expression in E. coli and Purification Previously, R. castenholzii nifH, nifB, nifD and
nifK were PCR-amplified, cloned and ligated into several different expression vectors with 6x
histidine tags: pACYC (NifH and NifB) and pETDuet1 (NifDK) (unpublished results) (Novagen,
Billerica, MA). Constructs encoding the nitrogenase homolog proteins NifH, NifB, and NifDK
solely and in strains for containing multiple constructs for co-expression have been transformed
into E. coli BL21 CodonPlus RIL (DE3) (Stratagene, Santa Clara, CA) cells for heterologous protein
expression. Cells were plated on Luria-Bertani (LB) agar plates with the appropriate antibiotics
(NifH and NifB- kanamycin (30μg/mL) and chloramphenicol (34μg/mL); NifDK- ampicillin
(100μg/mL) and chloramphenicol (34μg/mL)). Cell stocks were made by picking one colony from
agar plates and growing the colony overnight in 5mL of LB broth with the appropriate
antibiotics. 30μL of dimethyl sulfoxide (DMSO) was added to 970mL of the 5mL culture and flash
frozen to make a cell stock. Starter cultures were made by inoculating 50mL of LB broth
containing the appropriate antibiotics with a loop of cell stock and growing overnight at 37OC,
250RMP. Growths of 1L LB broth with amended antibiotics were inoculated with 5mL of starter
culture and cells were grown at 37OC, 250RPM until OD600 =0.5 when isopropyl β-D-1thiogalactopyranoside (IPTG) was added to a final concentration of 1mM. Ferrous ammonium
sulfate was also added to a final concentration of 191μM. After 2 hours of additional growth at
37OC, 250RPM, the cells were moved to 4OC, supplemented with an additional 191μM ferrous
17
ammonium sulfate and purged with nitrogen overnight. Cells were harvested by centrifugation
at 5000xg in an Avanti J-30I centrifuge (Beckman Coulter Inc., Brea, CA). Protein purification
facilitated by His-Tag nickel affinity chromatography was carried out under anaerobic conditions
(see below). Protein concentrations were determined using the Bradford assay with bovine
serum albumin (Sigma, St. Louis, MO) as the standard. Protein size and purity was monitored by
SDS-PAGE using a Precision Plus Protein Dual Color Ladder (BioRad, Hercules, CA) and western
blot using a his tag mouse monoclonal antibody (Lifespan Biosciences, Seattle, WA).
Roseiflexus castenholzii Nitrogenase
NifH Homolog Purification All buffers and reagents used were made anaerobic by
purging with nitrogen. Purification of the nitrogenase Fe protein homolog from Roseiflexus
castenholzii was completed by lysing the cells anaerobically using the following lysis buffer
(10mL per gram of cell pellet): 50mM HEPES pH8, 50mM NaCl, 10mM MgCl2, 1mM dithionite
(DT) 5% glycerol, 1% Triton X-100, 1mM phenylmethylsulfonyl fluoride (PMSF). The cells were
resuspended in lysis buffer, and lysed by rapid decompression using a “cell bomb” (Parr
Instrument Co., Moline, IL). The procedure for the cell bomb included pressurization for 1 hour
in under argon pressure at 1300psi followed by pressure release and removal of lysate from the
cell bomb through a pressure release valve. Following lysis, the lysate was cleared using
centrifugation for 30 minutes at 36000xg in an Avanti J-30I centrifuge (Beckman Coulter Inc.,
Brea, CA). Cleared lysate was then passed over a Ni2+-NTA affinity column (Thermo Scientific,
Waltham, MA) equilibrated with Buffer A. Buffer A had the following composition: 50mM TrisHCl pH 8.0, 500mM NaCl, 1mM DT, 5% glycerol. The Ni2+-NTA column was washed using 10
column volumes of wash buffer (50mM Tris-HCl pH 8.0, 500mM NaCl, 1mM DT, 10mM
imidazole, 5% glycerol). Protein was eluted from the column using 6 column volumes of elution
18
buffer (50mM Tris-HCl pH 8.0, 500mM NaCl, 1mM DT, 100mM imidazole, 5% glycerol). Protein
purity was monitored by SDS-PAGE and western blot using a his tag mouse monoclonal antibody
(lifespan Biosciences, Seattle, WA) and concentration was monitored spectrophotometrically by
Bradford assay [53], throughout the course of purification.
Roseiflexus castenholzii Nitrogenase
MoFe-homolog Protein Purification All buffers and reagents used were made
anaerobic by purging with nitrogen. Purification of the nitrogenase MoFe homolog from
Roseiflexus castenholzii was completed by lysing the cells anaerobically using the following lysis
buffer: 50mM HEPES pH8, 50mM NaCl, 10mM MgCl2, 1mM dithionite (DT) 5% glycerol, 1%
Triton X-100, 1mM PMSF. The cells were lysed in lysis buffer with constant stirring on a stir
plate. Following lysis, cellular debris was pelleted using centrifugation for 30 minutes at 36000xg
in an Avanti J-30I centrifuge (Beckman Coulter Inc., Brea, CA). The nifD and nifK gene products
were found to be present in the cell pellet. Following lysis, the cell pellet was resuspended in 8M
urea and centrifuged for 30 minutes at 36000xg. The supernatant was then added to a PD-10
desalting column packed with Sephadex G25 medium (GE Healthcare, Piscataway, NJ). Protein
was eluted using G25 buffer (50mM HEPES pH 7.4, 300mM NaCl, 5% glycerol). G25 elutions
were then passed over a Ni2+-NTA affinity column (Thermo Scientific, Waltham, MA) equilibrated
with Buffer B. Buffer B had the following composition: 50mM HEPES pH 7.4, 300mM NaCl, 1mM
DT, 5% glycerol. The Ni2+-NTA column was washed using 10 column volumes of wash buffer
(50mM Tris-HCl pH 8.0, 500mM NaCl, 1mM DT, 10mM imidazole, 5% glycerol). Protein was
eluted from the column using 6 column volumes of elution buffer (50mM Tris-HCl pH 8.0,
500mM NaCl, 1mM DT, 100mM imidazole, 5% glycerol). Protein purity was monitored by SDSPAGE and western blot analysis using a his tag mouse monoclonal antibody (Lifespan
19
Biosciences, Seattle, WA) and concentration was monitored spectrophotometrically by Bradford
assay [53] throughout the course of purification.
Roseiflexus castenholzii Nitrogenase
nifB Gene Product Purification All buffers and reagents used were made anaerobic by
purging with nitrogen. Attempts to purify the R. castenholzii nifB gene product were completed
by lysing the cells anaerobically using the following lysis buffer: 50mM HEPES pH8, 50mM NaCl,
10mM MgCl2, 1mM dithionite (DT) 5% glycerol, 1% Triton X-100, 1mM PMSF. The cells were
lysed in lysis buffer with constant stirring on a stir plate. Following lysis, cellular debris was
pelleted using centrifugation for 30 minutes at 36000xg in an Avanti J-30I centrifuge (Beckman
Coulter Inc., Brea, CA). Cleared lysate was passed over a Ni2+-NTA affinity column (Thermo
Scientific, Waltham, MA) equilibrated with Buffer A. The Ni2+-NTA column was washed using 10
column volumes of wash buffer (50mM Tris-HCl pH 8.0, 500mM NaCl, 1mM DT, 10mM
imidazole, 5% glycerol). Protein was eluted from the column using 6 column volumes of elution
buffer (50mM Tris-HCl pH 8.0, 500mM NaCl, 1mM DT, 100mM imidazole, 5% glycerol). Protein
purity was monitored by SDS-PAGE and western blot analysis using a his tag mouse monoclonal
antibody (Lifespan Biosciences, Seattle, WA) and concentration was monitored
spectrophotometrically by Bradford assay [53] throughout the course of purification.
Results and Discussion
Growth of Roseiflexus castenholzii
Without Fixed Nitrogen
In order to determine if R. castenholzii could grow without fixed N, it was grown in PE
media without fixed N and with supplemented alternative carbon sources: acetate, citrate,
glucose, glycyl-glycine and pyruvate. Growth curves of R. castenholzii with and without fixed N
20
indicated that growth without fixed N was less than growth with fixed N, by OD600. In growth
with fixed N the cultures repeatedly reached an OD600 of 0.6 within 48 hours at which time
growth plateaued (data not shown). In cultures without fixed N the maximum OD600 observed
was 0.3 in the presence of acetate. However, when the cultures without fixed N were
transferred in order to remove any remaining fixed nitrogen from the original inoculum, no
growth was observed (OD600 less than 0.04 monitored over 48 hours) (data not shown). These
results suggest that any growth observed without fixed N was due to remaining fixed N from the
original inoculum and R. castenholzii needs fixed N to grow under the conditions tested. Testing
the growth of R. castenholzii in media lacking fixed N and amended with other carbon sources
will further elucidate whether the organism can fix nitrogen. It would also be worthwhile to use
quantitative reverse-transcription PCR (qRT-PCR) to monitor the expression of nif-homolog
transcripts during growth with and without fixed N. If R. castenholzii is able to fix nitrogen, it
would be expected that the abundance of transcripts of the Nif protein homologs would
increase.
Expression and Purification of Roseiflexus
castenholzii Fe Protein Homolog
Initial results suggest NifH is being over-expressed in the E. coli host and can be purified
on a nickel affinity column (Figure 1.5) (western data not shown). The predicted size of the iron
protein homolog is 30.1kDa which was confirmed by SDS-PAGE (Figure 1.5) and western blot
(data not shown). The nifH gene product was found to easily precipitate in several different
buffers and preliminary iron assays revealed an iron number of zero from purified (nonreconstituted) protein (data not shown). Presence of a complete iron-sulfur cluster may be
necessary for formation of the NifH homodimer (which is bridged by the [4Fe-4S] cluster). The
21
protein was also found to precipitate quickly, suggesting the need for a different storage buffer.
In order to optimize protein solubility and increase yield, multiple buffered conditions were
tried, as well as different lysis procedures (including sonication, pressure cell, and
microfluidizing). Although the protein is expressed well and 0.4mg of protein was purified from
1g cell pellet, there are still contaminating proteins present (Figure 1.5), and a storage buffer in
which the protein would not precipitate was not found. Optimal conditions for purification and
Ladder
Elution 6
Elution 5
Elution 4
Elution 3
Elution 2
Elution 1
Wash 2
Wash 1
Pellet
storage of the Fe protein homolog are still being examined.
50kDa
37kDa
Figure 1.5. SDS-PAGE of purification of R. castenholzii
nitrogenase iron protein homolog.
Expression and Purification of Roseiflexus
castenholzii MoFe Protein Homolog
Initial results suggested that NifD and NifK are being over-expressed from a single plasmid
(petDUET) as a dimer in the E. coli host (Figure 1.6 (left)). The predicted sizes of NifD and NifK
Ladder
Soluble Fraction
Insoluble Fraction
Ladder
Soluble Fraction
Insoluble Fraction
22
50kDa
Figure 1.6. Initial purification of
NifD and NifK. On the left is an SDSPAGE gel. On the right is a western
blot of the SDS-PAGE gel.
are 53.0kDa and 52.8kDa which was confirmed by SDS-PAGE (Figure 1.6 (left)) and western blot
(Figure 1.6 (right)). Only NifK is his tagged, and therefore only one band is expected to be
present on a western blot. Further attempts at purification of the NifD and NifK proteins
revealed that the proteins were being expressed in inclusion bodies. Addition of a urea
precipitation step in the purification protocol resulted in removal of the proteins from the
inclusion bodies. Following urea precipitation, a desalting column and a Ni2+-NTA affinity column
were used to remove the urea and it was found that the proteins were no longer associated.
Figure 1.7 shows the G25 desalting column fractions with both NifK (52.80kDa) and NifD
23
Ladder
Elution 5
Elution 4
Elution 3
Elution 2
Elution 1
Ladder
Elution 6
Elution 5
Elution 4
Elution 3
Elution 2
Elution 1
(53.0kDa). The elutions from the G25 desalting column were added to a Ni2+-NTA affinity column
50kDa
Figure 1.7. Left: SDS-PAGE gel of G25 elution fractions of NifD
and NifK. Right: SDS-PAGE gel of Ni2+-NTA affinity column
elutions, showing only the presence of NifK.
where it was found that NifD and NifK were no longer associated, only one protein was present,
NifK (which is his tagged) as seen on an SDS-PAGE gel (Figure 1.7 (right)). NifK and NifD must be
associated in order to be eluted from the Ni2+-NTA column together because only NifK is his
tagged, and therefore only NifK will bind to the column if dimerization does not occur. The
identity of the protein in the elutions was confirmed to be NifK by western blot (data not
shown). These results suggest that after the urea precipitation step the NifD and NifK proteins
are not associating and are not eluted from the nickel affinity column bound together.
In order to favor the interaction of NifD and NifK following urea precipitation, the cleared
lysate was dialyzed into a series of different buffers varying in pH, salt concentration and
glycerol concentration. In all cases the protein precipitated almost immediately upon being
24
placed in the dialysis buffer (data not shown). A gradient of urea concentrations was also used in
order to see if less harsh conditions would release the proteins from the inclusion bodies and
facilitate re-folding. The only urea concentration which removed the proteins from the inclusion
bodies was 8M urea (data not shown). Previous research indicates pH of growth media and
temperature of induction significantly influence formation of inclusion bodies [54]. Therefore,
alterations to the growth procedures can be made in order to perhaps limit the formation of
inclusion bodies.
Expression and Purification of Roseiflexus
castenholzii NifB Protein Homolog
Initial attempts to overexpress NifB through heterologous expression in E. coli followed by
purification attempts suggested that NifB is not being over-expressed in the E. coli host (data
not shown). The predicted size of NifB is 34.3kDa, no product of that size was seen in any of the
purification fractions by SDS-PAGE or western blot when attempts at purification were made
(data not shown). Alternative growth conditions should be tried including altering growth
temperature and induction temperature. It is possible that a lower growth temperature would
allow the E. coli to grow more slowly, with plenty of time to produce the protein.
In summary, the results presented here suggest that R. castenholzii NifH and NifDK
homologs can be heterologously expressed in E. coli. Although purification protocols presented
herein did not produce soluble protein in quantities sufficient for crystallization trials, the results
did reveal several promising experiments to be tried in future studies that are likely to produce
soluble proteins.
25
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cluster assembly: the SUF machinery. Journal of Biological Inorganic Chemistry 10(7): p.
713-21.
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Muraki, N., J. Nomata, K. Ebata, T. Mizoguchi, T. Shiba, H. Tamiaki, G. Kurisu, and Y.
Fujita. (2010) X-ray crystal structure of the light-independent protochlorophyllide
reductase. Nature 465(7294): p. 110-4.
43.
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Biology 154: p. 434-438.
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enzyme for bacteriochlorophyll biosynthesis: reconstitution of chlorophyllide a
reductase with purified X-protein (BchX) and YZ-protein (BchY-BchZ) from Rhodobacter
capsulatus. Journal of Biological Chemistry 281(21): p. 15021-8.
45.
Sarma, R., B.M. Barney, T.L. Hamilton, A. Jones, L.C. Seefeldt, and J.W. Peters. (2008)
Crystal structure of the L protein of Rhodobacter sphaeroides light-independent
protochlorophyllide reductase with MgADP bound: a homologue of the nitrogenase Fe
protein. Biochemistry 47(49): p. 13004-15.
46.
Blankenship, R.E. (1992) Origin and early evolution of photosynthesis. Photosynthesis
Research 33: p. 91-111.
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Bjorn, L.O. and Govindjee. (2009) The evolution of photosynthesis and chloroplasts.
Current Science 96(11): p. 1466-1474.
48.
Boyd, E.S., T.L. Hamilton, and J.W. Peters. (2011) An alternative path for the evolution of
biological nitrogen fixation. Frontiers in Microbiological Chemistry 2.
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49.
Collins, A.M., Y. Xin, and R.E. Blankenship. (2009) Pigment organization in the
photosynthetic apparatus of Roseiflexus castenholzii. Biochimica et Biophysica Acta
1787(8): p. 1050-6.
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Staples, C.R., S. Lahiri, J. Raymond, L. Von Herbulis, B. Mukhophadhyay, and R.E.
Blankenship. (2007) Expression and association of group IV nitrogenase NifD and NifH
homologs in the non-nitrogen-fixing archaeon Methanocaldococcus jannaschii. Journal
of Bacteriology 189(20): p. 7392-8.
51.
Hanada, S., S. Takaichi, K. Matsuura, and K. Nakamura. (2002) Roseiflexus castenholzii
gen. nov., sp. nov., a thermophilic, filamentous, photosynthetic bacterium that lacks
chlorosomes. International Journal of Systemic and Evolutionary Microbiology 52(Pt 1):
p. 187-93.
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Hanada, S., A. Hiraishi, K. Shimada, and K. Matsuura. (1995) Chloroflexus aggregans sp.
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gliding movement. International Journal of Systematic Bacteriology 45(4): p. 676-81.
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30
CHAPTER 2
EXPRESSION AND PURIFICATION OF THREE CYANOBACTERIAL TRICARBOXYLIC ACID CYCLE
PROTEINS
Introduction
The Tricarboxylic Acid Cycle Overview
The tricarboxylic acid (TCA) cycle is a universal pathway used for energy generation by
all aerobic organisms [3]. Also known as the Kreb’s cycle or the citric acid cycle, this cycle is
responsible for using acetyl coenzyme A (acetyl-CoA) from glycolysis and amino acid and fatty
acid degradation to generate CO2, NADH, guanisine triphosphate (GTP) and reduced ubiquinone
(QH2) [4]. Although several metabolic pathways feed acetyl-CoA into the TCA cycle, the primary
source of acetyl-CoA is glycolysis, the set of enzymatic reactions that oxidizes one glucose
molecule to two pyruvate molecules. The products from the TCA cycle are used in the
biosynthesis of compounds such as amino acids and carbohydrates. The net reaction of the TCA
cycle is the following:
3NAD+ + FAD + GDP + Pi + acetyl-CoA + 2 H20 → 3 NADH + FADH2 + GTP + CoA + 2 CO2 + 3H+
This overall reaction is completed by a series of eight reactions catalyzed by eight different
enzymes [4]. In each round of the cycle one molecule of acetyl-CoA reacts with oxaloacetate in a
condensation reaction catalyzed by citrate synthase to form citrate. Citrate continues through
the cycle, two CO2 molecules are evolved and after eight reactions another molecule of
oxaloacetate is formed which can be used in another round of the cycle. Figure 2.1 shows how
oxaloacetate is regenerated in each turn of the cycle through the actions of the eight TCA
enzymes. The primary source of acetyl-CoA is glycolysis, which generates 2 molecules of acetyl-
31
Acetyl-CoA
Citrate
Synthase
Citrate
Oxaloacetate
Malate
Dehydrogenase
Malate
Aconitase
Isocitrate
Isocitrate
Dehydrogenase
α-ketoglutarate
Fumarase
Fumarate
Succinate
Dehydrogenase Succinate
α-Ketoglutarate
Dehydrogenase Complex
(2-Oxoglutarate
Dehydrogenase Complex)
Succinyl
-CoA
Succinyl-CoA
Synthetase
Figure 2.1 The Tricarboxylic Acid (TCA) Cycle. Blue boxes contain intermediates,
black lettering (without a blue box) denotes enzyme names.
CoA. Regeneration of oxaloacetate is important because it is needed to react with each acetylCoA. As mentioned previously, several metabolic pathways feed into the TCA cycle, which has
been called one of the most important metabolic pathways [5, 6]. Figure 2.2 illustrates the
importance of the TCA cycle because of its role as a “checkpoint” for numerous metabolites.
32
Proteins
Carbohydrates
Glucose-6Phosphate
Glycogen
Urea Cycle
Fats and Lipids
Lipogenesis
Pyruvic
Acid
Fatty Acid
Degradation
AcetylCoA
The
Tricarboxylic
Acid Cycle
Electron Transport
Chain
Figure 2.2. The interactions of several metabolic pathways, emphasizing the central
role of the tricarboxylic acid cycle in metabolism.
The Tricarboxylic Acid Cycle in Eukaryotes
In eukaryotes the TCA cycle takes place in the mitochondria where all of the enzymes
are located. The compartmentalization of the eukaryotic cell means that any reactants must be
transported to the mitochondria before they can be used in the cycle; similarly, any products
must be transported from the mitochondria to be used in other metabolic pathways. Several
transport proteins exist which actively transport pyruvate into the mitochondria where it can be
33
oxidized to form acetyl-CoA by the pyruvate dehydrogenase complex. The protein transporters
as well as all of the enzymes involved in the TCA cycle in eukaryotes have been extensively
studied. Each enzyme in the cycle is required; when one enzyme is missing or non-functional
metabolic disorders which are often encephalomyopathies result [5-8]. The importance of each
enzyme in the cycle has been illustrated by the presence of humans with mutations or deletions
in the genes encoding TCA enzymes [9-11]. Also, mouse knockout or knockdown models in
which a gene coding for a TCA enzyme is inactivated or less functional have allowed scientists to
thoroughly investigate the role of each enzyme in the TCA cycle. The results of the mouse
genetic studies have been invaluable in studying metabolic disorders of the TCA cycle and
highlighting the importance of each enzyme in the cycle [12-14].
The Tricarboxylic Acid Cycle in Prokaryotes
The TCA cycle has the same functions in prokaryotes as in eukaryotes: oxidize two
carbon units from acetyl-CoA to CO2 and provide precursors to be used in metabolic synthesis.
The steps of the cycle are the same in aerobic prokaryotes and eukarya, with a few exceptions.
Prokaryotes lack membrane-bound organelles, so the TCA cycle takes place in the cytoplasmic
matrix. The complete 8 enzyme TCA cycle is present in many bacterial species, however there
are also species that use an abbreviated version of the TCA cycle called the glyoxylate cycle or
glyoxylate shunt (Figure 2.3) [15]. The two reactions that are unique to the glyoxylate pathway
are:
Isocitrate → succinate + glyoxylate (catalyzed by isocitrate lyase)
Glyoxylate + acetyl-CoA → malate + CoA (catalyzed by malate synthase)
34
Acetyl-CoA
Oxaloacetate
Malate
Citrate
Acetyl-CoA
Fumarate
Isocitrate
Glyoxylate
+
Succinate
Figure 2.3. The glyoxylate cycle. Steps that differ from the TCA cycle are
shown in light pink.
The glyoxylate cycle is useful because it regenerates the TCA cycle intermediates malate and
succinate, bypassing the reactions that release carbon as CO2 [3, 16, 17]. When carbohydrate
availability is low, the glyoxylate pathway can be used to make more malate, a precursor of
oxaloacetate, which can eventually be used in gluconeogenesis. Many different bacteria such as
E. coli and A. vinelandii are known to use the glyoxylate cycle [18, 19]. A. vinelandii primarily
35
uses the glyoxylate cycle to metabolize acetate and when grown in the presence of both glucose
and acetate the organism exhibits a diauxic growth curve in which growth occurs in two phases
separated by a lag phase. In the first phase of growth the bacteria is metabolizing sugar of
preference (glucose), then growth slows while the machinery (glyoxylate shunt enzymes)
needed to metabolize the second sugar is made and finally log growth resumes on the second
carbon source.
In other prokaryotes such as Mycobacterium tuberculosis (Mtb), the TCA cycle is
completed using alternatives to some of the enzymes. In Mtb a thiamine pyrophosphate
dependent enzyme called 2-oxoglutarate decarboxylase (2-OGDC) produces succinic
semialdehyde which is oxidized by succinic semialdehyde dehydrogenase (SSADH) to form
succinate; these reactions bypass the reactions catalyzed by 2-oxoglutarate dehydrogenase (also
known as α-ketoglutarate dehydrogenase) which is a rate limiting step in the TCA cycle, and
succinyl Co-A synthetase (Figure2.1)[20]. 2-OGDC converts α-ketoglutarate to succinic
semialdehyde which is then oxidized to succinate by succinic semialdehyde dehydrogenase
(SSADH) [20]. Recently, a similar alternative TCA pathway was elucidated in a group of
photosynthetic bacteria, the cyanobacteria [21-23].
There are three different groups of gram-negative photosynthetic bacteria: purple
bacteria, green bacteria, and cyanobacteria. Purple bacteria are named because they contain
bacteriochlorophyll a or b and carotenoids, pigments which make them appear anywhere from
purple to orange [24-26]. Green bacteria contain a different set of bacteriochlorophylls and
carotenoids which make them green [27]. Both purple and green photosynthetic bacteria have
only photosystem I and are therefore able to grow using anoxygenic photosynthesis [15]. Due to
the inability of purple and green bacteria to use water as an electron source, they typically
36
occupy deep anoxic zones where hydrogen sulfide, sulfur and hydrogen are available as electron
donors [28]. In contrast, cyanobacteria use chlorophyll a and photosystems I and II which allow
them to complete oxygenic photosynthesis.
Cyanobacteria are photolithoautotrophs (with few exceptions) that represent a very
large, extremely diverse phylum of bacteria that has been of interest to evolutionary scientists
because of their widespread presence and influence on early Earth’s development.
Cyanobacterial fossils are some of the oldest fossils ever found, dating back 3.5 billion years
[29]. It is believed that cyanobacteria and methanogenic bacteria were largely responsible for
the development of Earth’s atmosphere; cyanobacteria produced O2 through photosynthesis
and methanogens produced methane (CH4) [30, 31]. Together, these gases contributed to the
change from a reducing atmosphere to an oxidizing atmosphere. Eventually the changes in the
atmosphere made the environment more hospitable for life [30-32]. In addition to the role
cyanobacteria are hypothesized to have played in the development of Earth’s atmosphere, they
are of interest because unlike many fossilized organisms, extant relatives of fossilized
cyanobacteria are widespread on Earth today, which allows scientists to use them in
evolutionary studies [33]. As organisms that use photosynthesis and the TCA cycle, extant
cyanobacteria provide invaluable insight into the evolution of both metabolic processes.
Photosynthesis particularly has had a profound impact on increasing the oxygen concentration
of the atmosphere to current levels. Phylogenetic studies place even more importance on
cyanobacteria, because the chloroplasts of plants and algae are believed to be derived from a
cyanobacterial ancestor; this also implies that cyanobacteria played a role in Eukaryotic
evolution [34]. Because of the evolutionary insight that can be gained from studying
37
cyanobacteria, they have been the focus of countless studies on the evolution of
photosynthesis, N2 fixation and the TCA cycle [21, 23, 29, 35, 36].
Although the classification of cyanobacteria is still debated, it is widely accepted that
cyanobacteria differ in their mode of reproduction, morphology, size, trichome arrangement
and ability to fix nitrogen. Many cyanobacteria are phototrophic facultative anaerobes, able to
switch from oxygenic photosynthesis to anoxygenic photosynthesis when conditions change
[35]. Despite the many differences between the classes of cyanobacteria, until recently it has
been widely accepted that cyanobacteria have an incomplete TCA cycle [21, 23, 36, 37]. Two
enzymes in the TCA cycle, α-ketoglutarate dehydrogenase (also known as 2-oxoglutarate
dehydrogenase) and succinyl-Co-A synthetase are not found in cyanobacteria. It was therefore
assumed that cyanobacteria were employing the pentose phosphate pathway to regenerate
NADPH and other intermediates for use in metabolic biosynthesis [21, 38].
In a recent study of electron flow in the metabolism of Synechocystis spp. Strain PCC
6803 it was found that when 2-oxoglutarate (also known as α-ketoglutarate) is added to media
an increase in succinate concentration is observed [39]. This study suggests the 2-oxoglutarate is
being converted to succinate by enzymes other than the traditional TCA enzymes 2-oxoglutarate
Figure 2.4. Conversion of α-ketoglutarate to succinate.
38
dehydrogenase and succinyl-CoA synthetase. It was recently discovered through genomic and
molecular studies of Synechococcus sp. PCC 7002 that genes encoding novel 2-oxoglutarate
decarboxylase (2-OGDC) and succinic semialdehyde dehydrogenase (SSADH) enzymes are
present [23]. These enzymes convert 2-oxoglutarate to succinate, bypassing the reactions
catalyzed by 2-oxoglutarate dehydrogenase (2-OGDH) and succinyl Co-A synthetase (Figure 2.4).
In the same report, it was also found that genes for these novel TCA enzymes are found in all
cyanobacterial genomes except those of Prochlorococcus and marine Synechococcus species
(Figure 2.5). The finding that succinate is produced in cultures of Synechocystis sp., in
combination with the recent discovery of two novel alternative enzymes involved in the TCA
cycle of Synechococcus strongly suggest that the cyanobacterial TCA cycle is not incomplete, as
has been believed for many years.
Zhang and Bryant completed preliminary mutational and biochemical studies on the
novel 2-OGDC and SSADH [23]. Several mutant Synechococcus strains which overexpressed one
or both of the novel enzymes displayed altered growth curves, indicating the metabolic role of
each enzyme. It was found that during growth in constant light, overexpression of 2-OGDC
slowed growth considerably, deletion mutants of both 2-OGDC and SSADH grew slower than the
wildtype strain (by 17%) and strains overexpressing SSADH grew slower than the wildtype as
well (but with less of a significant decrease in growth than was observed during overexpression
of 2-OGDC). 2-oxoglutarate is a very important metabolite for ammonia assimilation, regulation
of nitrogen and carbon metabolism and as a precursor for heme and chlorophyll biosynthesis
[40]. It was hypothesized by Zhang and Bryant that growth rates are slowed in the mutants
overexpressing one or both of the novel TCA enzymes because of the importance of 2oxoglutarate in metabolism and the detrimental effects that depleting it could have [40].
39
Citrate
Oxaloacetate
Isocitrate
2Oxoglutarate
Malate
2-Oxoglutarate
Dehydrogenase
Succinic
Semiadehyde
Fumarate
Succinate
Succinic
Semialdehyde
Dehydrogenase
Figure 2.5. Proposed alternative to the TCA cycle in the cyanobacteria,
Synechococcus sp. PCC 7002.
The importance of the novel cyanobacterial 2-OGDC and SSADH enzymes in the TCA
cycle is evident because it provides new evidence on the completeness of the TCA cycle in
cyanobacteria which could also lead to new insights into evolution of the TCA cycle from
cyanobacterial ancestors to extant cyanobacteria. Additional biochemical studies were
completed on a recently identified fumarase (designated A2041) from Synechococcus sp. 7002
[41, 42]. Interestingly, biochemical studies have found that this fumarase catalyzes the
reversible reaction between fumarate and malate and performs the half reactions at different
optimal pH values [42]. This suggests a novel reaction mechanism for this fumarase [42]. This
project was designed in order to further characterize the novel fumarase, 2-OGDC and SSADH
enzymes from Synechococcus sp. PCC 7002 by determining their structure through x-ray
crystallography.
40
Materials and Methods
Growth of Select
Synechococcus sp. PCC 7002 TCA Enzymes
Previously, the open reading frames SynPCC7002_A2041, SynPCC7002_A2770 and
SynPCC7002_A2771 (encoding the fumarase, 2-OGDC and SSADH, respectively) were separately
amplified by polymerase chain reaction (PCR) and cloned and ligated into expression vector
pAQ1 with a 10x histidine tag [23, 43]. E. coli NEB5-alpha cells (NEB, Ipswich, MA) were
transformed with pAQ1 encoding SynPCC7002_A2041, SynPCC7002_A2770 or
SynPCC7002_A2771. The pAQ1 plasmid was modified to contain resistance genes for gentamicin
(when SynPCC7002_A2770 was the insert) or spectinomycin (when SynPCC7002_A2041 or
SynPCC7002_A2771 was the insert) [43]. Following transformation, cells were plated on LuriaBertani (LB) agar plates with ampicillin (100μg/mL) and either gentamicin (20μg/mL) or
spectinomycin (20μg/mL) for selective growth of the respective plasmids. Single colonies were
picked from the agar plates and grown in 5 mL of LB broth with the appropriate antibiotics.
From the 5mL growths, cell stocks of each plasmid were made into by adding 970μL overnight
culture to 30μL DMSO, after which they were flash frozen in liquid nitrogen and stored at -80OC
until use.
For protein expression, 1 loop of cell stock was added to 50mL LB broth amended with
the appropriate antibiotics (see above) and grown for 8 hours at 37OC, 250 RPM. At an OD600 of
approximately 0.8, 5mL of starter culture were added to 1L of LB broth amended with the
appropriate antibiotics. The 1L culture was grown overnight (~12hours) at 37OC, 250RPM, the
protein was expressed constitutively; expression was not under control of the T7 promoter.
Following growth, the cells were harvested by centrifugation at 4OC, 5,180xG in an Avanti J-30I
41
centrifuge (Beckman Coulter Inc., Brea, CA), weighed, flash frozen in liquid nitrogen and stored
at -80OC until use.
Purification of Select
Synechococcus sp. PCC 7002 TCA Enzymes
Cells were lysed in lysis buffer (10mL/g cell pellet) (50mM Tris-HCl pH 8.0, 50mM NaCl,
5% w/v glycerol, 1% triton X-100, 100mM MgCl2 and 1mM phenylmethylsulfonyl fluoride
(PMSF)) by constant stirring on a stir plate at room temperature. Following lysis, soluble lysates
were obtained by centrifugation at 36000Xg and loaded on Ni2+-NTA affinity resin (Thermo
Scientific, Waltham, MA) equilibrated with 50mM Tris-HCl pH 8.0, 50mM NaCl, and 100mM
MgCl2. After the soluble lysates had flowed through the affinity column, the column was
washed with wash buffer (10mM imidazole, 300mM NaCl and 50mM Tris-HCl pH 8.0) and
proteins were eluted with elution buffer (300mM imidazole, 300mM NaCl, 50mM Tris-HCl pH8).
Fractions containing the recombinant proteins were monitored using sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE) and size was monitored using a Precision Plus
Protein Dual Color ladder (BioRad, Hercules, CA). Protein-containing elutions were concentrated
using Amicon Ultra (10,000Da MWCO) filters (Millipore, Billerica, MA) to no more than 10mg/mL
by centrifugation at 5000xg. Concentration of protein was monitored using the Bradford assay
[44]. Following concentration, protein was further purified by size exclusion chromatography
using a Sephacryl S-200, 1.5cm x 80cm column (Amersham Biosciences/Ge Healthcare,
Piscataway, NJ) equilibrated with running buffer (300mM NaCl, 20mM Tris pH8.0, 5% glycerol)
and calibrated with gel filtration standards (BioRad, Hercules, CA) at a flow rate of 0.5mL/min.
Protein purity was monitored by SDS-PAGE and western blot using a His tag mouse monoclonal
antibody (Lifespan biosciences, Seattle, WA) and concentration was monitored using the
42
Bradford Assay [44]. Protein fractions without contaminating proteins, determined by SDSPAGE, were concentrated using Amicon Ultra filters (Millipore, Billerica, MA) to 15mg/mL.
Hanging drop crystallization screens with 1μL protein (15mg/mL) and 1μL crystallization
solution were completed using Emerald Cryo I and II crystallization solutions and Hampton
Crystal Screen Cryo™ and Natrix™ crystallization solutions (Emerald Biosciences, Bainbridge
Island, WA and Hampton Research, Aliso Viejo, CA). To rule out salt crystals, any crystals that
grew were tested with Izit Crystal Dye™ (Hampton Research, Aliso Viejo, CA). Fumarase crystal
screens were completed using a Honeybee Nanodrop robot (Genomic Solutions Ltd., Ann Arbor,
MI) using 96-well sitting drop trays and Hampton crystallization solutions in the following
crystallization screens: Index™, Salt Rx™, Peg Rx™, Peg/ion™, Crystal Screen Light™, Crystal
Screen Cryo™ (Hampton Research, Aliso Viejo, CA). An additional 96-well sitting drop tray was
made using the robot and the Emerald Cryo Screen™ crystallization solutions (Emerald
Biosciences, Bainbridge Island, WA).
Results and Discussion
Fumarase
A plasmid containing SynPCC7002_A2041 (the novel fumarase) from Synechococcus sp.
PCC 7002 was transformed into E. coli NEB5α cells which were then grown in LB media,
harvested and the fumarase protein was purified using a Ni2+-NTA affinity column and a size
exclusion column. Growth of transformed NEB5α cells overnight in 1L LB media yielded a cell
pellet of approximately 3g and 0.5mg of purified protein per gram cell pellet. Following flash
freezing, protein was purified by lysing the cells in a lysis buffer (see above) and lysis was
Ladder
Elution 8
Elution 7
Elution 6
Elution 5
Elution 4
Elution 3
Elution 2
Elution 1
Wash 2
Wash 1
Flow Through
Cleared Lysate
pellet
lysate
43
50kDa
Ladder
Elution 6
Elution 5
Elution 4
Elution 3
Elution 2
Elution 1
Figure 2.6. SDS-PAGE gel of Fumarase purification. Elutions are from
the Ni2+-NTA affinity column.
50kDa
Figure 2.7. Western blot with
His-tag antibody of fumarase
Ni2+-NTA elutions 1-6.
44
monitored by SDS-PAGE (Figure 2.6). Figure 2.6 shows fractions from a purification of fumarase
in which there is a distinct band in all of the elutions at just less than 50kDa. The calculated size
of the fumarase protein is 54.6kDa, larger than the band seen on the gel (Figure 2.6). However,
analysis by western and SEC indicated that the purified protein is the fumarase protein, but it
runs anomalously on SDS-PAGE gels (Figures 2.6 and 2.7). Previous studies of fumarase proteins
indicate that the functional protein is a tetramer [45, 46]. The results from size exclusion
chromatography confirm that the novel fumarase from Synechococcus sp. PCC 7002 is also a
tetramer (Figure 2.8).
Figure2.8. SEC peak of Fumarase. The enzyme is present as a tetramer, as
calculated from a standard curve.
45
Preliminary crystallization screens using the Honeybee Nanodrop robot (Genomic
Solutions Ltd., Ann Arbor, MI) showed no crystal growth. Future studies include optimizing the
buffer in order to maintain protein stability while concentrating beyond 15mg/mL.
Succinic Semialdehyde Dehydrogenase
A plasmid containing the novel SSADH (SynPCC7002_A2771) from Synechococcus sp.
PCC 7002 was transformed into E. coli NEB5α cells which were then grown in LB media,
harvested and the SSADH protein was purified using a Ni2+-NTA affinity column and a size
exclusion column (see above). Growth of transformed NEB5α cells overnight in 1L LB media
yielded a cell pellet of approximately 2.5g and 0.6mg of pure protein per gram of cell pellet.
Following flash freezing, protein was purified by lysing the cells in a lysis buffer (see above) and
lysis was monitored by SDS-PAGE (Figure 2.9). Figure 2.9 shows fractions from a purification of
SSADH in which there is a distinct band in all of the elutions at approximately 50kDa. The
predicted size of SSADH is 51.1kDa, which is consistent with the size seen by SDS-PAGE and
confirmed by western blot (Figures 2.9 and 2.10). Previous research on SSADH proteins from
other organisms indicates they are tetramers, which is consistent with what was seen in this
project by size exclusion chromatography (Figure 2.11)
Ladder
Elution 6
Elution 5
Elution 4
Elution 3
Elution 2
Elution 1
Wash 2
Wash 1
Through
Flow
Lysate
Cleared
Pellet
Lysate
46
50kDa
Elution 1
Elution 2
Elution 3
Elution 4
Elution 5
Elution 6
Ladder
Figure 2.9. Purification of SSADH. Elutions are from the Ni2+-NTA affinity
column.
50kDa
Figure 2.10.Western blot
using His-tag antibody of
SSADH Ni2+-NTA elutions 16.
47
Figure 2.11. SEC peak of SSADH. The enzyme is present as a tetramer, as calculated
from a standard curve.
[47]. Analysis by western also confirmed that the histidine-tagged SSADH was present in the
elutions (Figure 2.11).
Preliminary crystallization screens using crystallization solutions from Hampton
Research and Emerald Biosciences resulted in no protein crystal growth. The next step in trying
to crystallize this protein is to change the salt conditions (both the concentration and the type of
salt) of the current buffer the protein is in (currently 300mM NaCl), as well as the concentration
of the protein, then setting up additional crystallization trays. A lower salt concentration may be
useful in helping to limit the formation of salt crystals; also, trying a different salt such as
potassium chloride may reduce salt crystal formation.
48
2-Oxoglutarate Decarboxylase
A plasmid containing the novel 2-OGDC (SynPCC7002_A2770) from Synechococcus sp.
PCC 7002 was transformed into E. coli NEB5α cells which were then grown in LB media,
harvested and the 2-OGDC protein was purified using a Ni2+-NTA affinity column and a size
exclusion column (see above). Growth of transformed NEB5α cells overnight in 1L LB media
yielded a cell pellet of approximately 3g. Following flash freezing, protein was purified by lysing
the cells in a lysis buffer (see above) and lysis was monitored by SDS-PAGE (Figure 2.12). Figure
2.12 shows fractions from a purification of 2-OGDC in which there is a faint band in all of the
elutions at approximately 55kDa. The predicted size of 2-OGDC is 56.9kDa, which is consistent
with the size observed by SDS-PAGE (Figure 2.12). Analysis by western blot also confirmed that
the histidine-tagged 2-OGDC was present in the elutions (Figure 2.13). However, protein
concentrations were too low for purification and subsequent crystallization screens.
Ladder
Elution 6
Elution 5
Elution 4
Elution 3
Elution 2
Elution 1
Wash 2
Wash 1
Flow Through
Cleared Lysate
Pellet
Optimization of the expression of 2-OGDC including growing the E. coli at a lower temperature,
50kDa
Figure 2.12. SDS-PAGE gel of 2-OGDC lysis. Elutions are
from Ni2+-NTA column.
Elution 6
Elution 5
Elution 4
Elution 3
Elution 2
Elution 1
Ladder
49
50kDa
Figure 2.13. Western blot of 2-OGDC
Ni2+-NTA affinity column elutions.
altering the pH of the growth media, transformation of the A2770 gene into an alternative
plasmid, and making different constructs of the protein should help increase the homologous
expression of the protein. Lowering the growth temperature of the E. coli will allow the bacteria
to grow more slowly, giving more time for protein expression. Changing the pH of the media and
altering the protein construct are both alterations of the growth protocol which have been
previously shown to increase protein expression of other difficult to express proteins[48] .
In summary, the results shown here indicate the ability to express and purify both the
novel fumarase and the novel SSADH from Synechococcus sp. PCC 7002 at concentrations
sufficient for crystallization screens. Additional crystallization conditions need to be used for
both the fumarase and the SSADH in order to grow crystals for x-ray crystallization studies. Work
completed on the novel 2-OGDC indicates the need for alternative expression or growth
conditions to increase the yield of protein.
50
References
1.
TCA/Kreb's/Citric Acid Cycle. 2011 [cited March, 2012]; Available from:
http://www.exrx.net/ExInfo/MetabolicDiagrams/TCAKrebsCitricAcidCycle.html.
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54
CHAPTER 3
ECOLOGICAL SIGNIFICANCE OF A SULFATE RESPIRING MICROORGANISM FROM AN ACIDSULFATE-CHLORIDE GEOTHERMAL FEATURE IN YELLOWSTONE NATIONAL PARK
Introduction
The Mercury Cycle
Mercury (Hg) is present in a wide variety of terrestrial, marine and freshwater
environments and is a known neurotoxin which cannot be detoxified by most higher forms of
life [3-6]. There are three common oxidation states of Hg: elemental (Hg0), mercurous (Hg+1 or
HgI) and mercuric (Hg+2 or HgII). Mercuric mercury is the most common form of Hg and is present
in natural systems as both inorganic and organic chemical compounds [3]. The predominant
inorganic HgII compounds in natural systems exist as ions complexed with selenium (HgSe),
oxygen (HgO), chloride (HgCl2), whereas a number of organomurcury compounds have been
identified that typically consist of linear methyl chains [e.g., methylmercury (CH3Hg+)].
Methylmercury (denoted here as MeHg), which will be discussed in more detail below, is
extremely toxic to higher forms of life due to its high affinity for thiol groups. Sources of Hg in
the environment include both natural (volcanoes, soil and water deposits) and anthropogenic
(coal burning power stations, municipal incinerators and recycled automobiles) sources [2, [7-9],
with natural sources considered to be the predominant source of Hg in the environment prior to
the industrial revolution [10]. Indeed, the evolution of biological mechanisms to cope with Hg
toxicity appears to be old, and is most likely to have originated over 2.5 billion years ago in a
hydrothermal environment; these environments are typified by high flux of natural Hg [10].
55
Microorganisms employ five known mechanisms to avoid Hg toxicity [11]: 1). reduced
uptake of inorganic mercuric ions, 2). reductive demethylation of MeHg followed reduction of
HgII to volatile Hg0 (mer encoded functions) which can then diffuse out of the system, 3).
sequestration of MeHg, 4). inorganic Hg methylation and 5). enzymatic reduction (mer encoded
functions) of inorganic HgII to Hg0 [11]. The first mechanism of Hg resistance, reduced uptake of
mercuric ions, has been characterized in a strain of Enterobacter aerogenes that uses two
plasmid encoded proteins to reduce cellular permeability to Hg2+ ions [12]. Demethylation of
MeHg and conversion to mercuric sulfide compounds was first noted in Clostridium cochlearium
T-2P [13]. This organism was found to have two plasmid-encoded genetic factors that react with
and inactivate MeHg to form insoluble mercuric sulfide. Desulfovibrio desulfuricans API
sequesters MeHg by producing hydrogen sulfide as a product of sulfate reduction, which is then
available to react with MeHg forming insoluble dimethylmercury sulfide [14]. Mercury
methylation is the subject of intense study due to the fact that MeHg is the most toxic form of
Hg for most organisms. However, to some bacteria such as Desulfovibrio desulfuricans LS, MeHg
is a less toxic form of Hg possibly due to subsequent sequestration or volatilization from the cell
[15]. The fifth form of Hg protection is enzymatic reduction, found in gram positive and gram
negative bacteria as well as some archaea. Enzymatic reduction of Hg is the result of the activity
of the gene products from the mer operon. The mer genes and their products have been widely
characterized and the biochemistry of how mer gene products function has been examined in
great detail [16-21]. Generally speaking, the mer system of resistance works by HgII being
sequestered (in inorganic or organic forms) and brought into the cell by transport proteins
where it is reduced by enzymatic detoxification to gaseous Hg0 and can diffuse out of the cell.
56
Two types of Hg resistance are encoded by the mer operon in bacteria: broad and
narrow [22]. In broad resistance, organisms are resistant to both inorganic and organic forms of
Hg. Typically these organisms have complex operons encoding for a suite of functions (e.g.,
merRTPAGBDE) [22]. Narrow resistance is characterized by only being resistant to inorganic
forms of Hg which can be detoxified by the actions of the merA gene product (discussed below)
[22]. A typical gene cluster which would provide narrow spectrum resistance to Hg would
contain at least merRTPADE genes [22]. The merR and merD gene products are involved in
detection of HgII; merR is also the regulatory protein responsible for turning on or off the
expression of the mer operon. merT and merP are involved in transport and mobilization of HgII
[11]. Enzymatic detoxification occurs through the actions of merA and B gene products which
are a mercury reductase and a organo-mercury lyase, respectively [23]. Hg is transported by the
MerE protein, a broad Hg transporter protein [24]. MerG is involved in phenylmercury
(C6H5Hg+)resistance by reducing cell permeability to this compound [25].
Through the actions of the mer operon gene products, organic forms of Hg such as
MeHg are reduced to Hg0. The mer operon is widely distributed in nature, and the presence of
merA genes in early branching thermophilic bacteria and archaea suggests that the origin of the
mer operon is from a hydrothermal environment where Hg levels have been naturally elevated
for billions of years [10]. In order to inhabit hydrothermal environments with high levels of Hg,
resistance mechanisms, such as mer operon encoded resistance must have evolved early, not in
the several hundred years since the industrial revolution increased levels of Hg in some
environments [10]. It is therefore likely that the most primitive Hg resistance mechanisms can
be identified and studied by examining organisms inhabiting thermal environments [10]. In
addition to mer encoded resistance mechanisms, the resistance mechanism of Hg methylation is
57
also likely to have evolved in a thermal area. By studying organisms in thermal environments, it
is therefore likely that the organisms are ancestors of the organisms encoding the earliest forms
of these resistance mechanisms.
In addition to biotic sources of MeHg, Hg can be methylated by abiotic mechanisms.
Abiotic MeHg is produced by transmethylation from fulvic and humic acids, and nonenzymatic
methylation of HgII by methylcobalamin, methyllead and methyltin and reaction of HgII with
small organic molecules such as methyliodide and dimethylsulfide [8, 26]. Abiotic methylation of
Hg can proceed through carbocationic, marbanionic or radical methyl groups depending on the
chemical properties of the original metal component of the methylating agent [27]. The rates at
which these abiotic reactions occur differ based on several environmental factors including pH,
the methyl donor that is present, temperature and presence of chelating agents in solution [8].
For example, in a freshwater environment where methylcobalamin is the methyl donor, the
methylation of Hg is likely to occur at faster rates if the environment is also acidic with low ionic
strength and low chloride levels [8]. In contrast, in a sea water environment where methyltin is
typically the methyl donor, the methylation reaction will occur more quickly if the environment
is alkaline with high chloride concentrations [8].
Biological MeHg is thought to be produced primarily by two guilds of organisms, the
sulfate-reducing bacteria and the iron-reducing bacteria [28]. Sulfate-reducing bacteria (SRB)
use sulfate (SO42-) as a terminal electron acceptor coupled with the oxidation of organic matter
or H2 to produce hydrogen sulfide (H2S) as shown in the following reactions in which CH2O
represents any organic compound that is oxidized:
2H+ + SO42- + CH2O→ H2S + 2CO2 + 2H2O
SO42- + 5H2→ H2S + 4H2O
58
Iron-reducing bacteria perform a similar reaction using Fe3+ rather than SO42- [29]. These two
groups of microorganisms are generally thought to be the primary sources of biotic MeHg in
natural systems [30-34]. SRB have been extensively studied, and for a long time were thought to
be the only microorganisms capable of producing MeHg [35]. When it was discovered that iron
reducing bacteria (IRB) also produced MeHg, for many years they were found to produce MeHg
at much slower rates than SRB [28, 36]. Only recently have IRB isolates been found that can
produce MeHg at rates equivalent to SRB (Figure 3.1) [28, 37].
Hg(0)
Hg(II)
SO42-→S2SRB
IRB
3+
Fe → Fe2+
CH3Hg
Figure 3.1 Mercury metabolism by
microorganisms; Hg methylation by
sulfate and iron reducing organisms
shown in yellow.
While MeHg is not toxic to some microorganisms, it is one of the most toxic forms of Hg
for higher eukaryotes due to its high affinity for thiol groups which contributes to its ability to
form a complex with the amino acid cysteine and the lipid solubility of that complex [5]. Higher
eukaryotes cannot process MeHg and as a result, it bioaccumulates in the food web [38].
Bioaccumulation can occur as follows: water-dwelling microorganisms (picophytoplankton)
containing MeHg tend to be consumed by invertebrates which are often a predominant food
source supporting fish and higher trophic levels such as birds. Because the invertebrates and
59
higher forms of life are incapable of reductively demethylating MeHg and each trophic level
needs to consume approximately 10-fold greater biomass than the previous trophic level, the
amount of MeHg found in the animals’ tissues tends to increase in the food chain. Samples of
birds’ feathers collected near environments with industrial Hg contamination or in areas of
naturally high levels of Hg have been found to have very high levels of MeHg, in some cases
nearly toxic levels [39, 40]. There have been several studies of MeHg levels in the feathers of
predatory birds which show elevated levels of MeHg [40, 41]. Likewise, humans which also
cannot process or detoxify MeHg, have the potential to bioaccumulate significant amounts of
MeHg through the consumption of fish, especially predatory fish at the top of the food chain
(e.g., tuna) [4].
In the body, MeHg quickly reacts with thiol groups, such as those present in the amino
acid cysteine [3]. The molecule formed from this reaction resembles the neutral amino acid
methionine; as a neutral molecule it can enter cells (Figure 3.2). Because it is smaller than other
organo-mercury species, mono-MeHg is more efficiently absorbed by animal cells [7]. Once
inside cells, MeHg often complexes with reduced glutathione (a common tripeptide that
functions as an antioxidant) [3]. Glutathione carrier proteins allow the MeHg-glutathione
CH3-Hg+ +
-
S-CH2-CH-COONH3+
CH3-Hg-S-CH2-CH-COONH3+
Cysteine/Hg Complex
CH3-S-CH2-CH2-CH-COONH3+
Methionine
Figure 3.2. The complex that forms between methylmercury
and cysteine and its resemblance to methionine.
60
complex to pass out of cells, travel through the body and even the blood brain barrier and be
deposited in tissues. Once in the tissues, (including the central nervous system), the MeHgglutathione complex causes neurologic symptoms to develop which include numbness of limbs,
loss of vision and hearing, ataxia, nausea and insomnia [3]. These affects are especially acute in
fetuses and infants due to the fact that their neurological systems are still in the developmental
stage.
Methyl Mercury in Yellowstone National Park
Geothermal systems, such as those in Yellowstone National Park, Wyoming, (YNP)
represent natural sources of Hg in the environment and are likely to harbor lineages of
organisms that have evolved in response to Hg stress for over 2.5 billion years. Given the
decreased ecosystem complexity in these environments, as well as the deeply branching nature
of many lineages that inhabit these environments, YNP springs have been a target for studies
aimed at understanding the evolutionary origin of Hg detoxification mechanisms. Soils and
geothermal springs in YNP have been shown to contain very high levels of Hg due to both past
as well as present day input from hydrothermal fluids that interact and which solubilize cinnabar
deposits in the subsurface [39, 42, 43]. Total Hg levels as well as MeHg levels are elevated in
environments in YNP. Hg and MeHg are concentrated in photosynthetic mats comprised of
Zygogonium, a species of green algae. Zygogonium mats are predominant in acidic, (pH 2-3.1)
moist surface environments near springs and seeps but not typically in flowing water [44]. This
species is noted for its purple vacuolar pigment and single, large chloroplast [45]. A thick
mucilaginous region (up to 5μm) is present exterior to the cell wall of Zygogonium, making it
resistant to large fluctuations in temperature and moisture [44, 46]. Zygogonium mats appear
reddish-purple in color due to an iron-tannin complex that accumulates in vacuoles within the
61
cells [47]. Studies of the Park ecosystem have demonstrated bioaccumulation of MeHg from the
stratiomyid (Diptera: Stratiomyidae) larvae that feed on Zygogonium algal mats with further
bioaccumulation occurring in birds that feed on these larvae [39]. Previously, it was shown that
larvae feeding on mats accumulate high levels of MeHg. MeHg is detectable in the gut and
tracheal tissue of the larvae. When a feather from a bird feeding on the larvae was collected and
analyzed for MeHg content it was found to contain six-fold more MeHg than the larval tissue
[39]. These results support bioaccumulation of MeHg through the food chain in YNP and
furthermore, suggest that Zygogonium microbial mats are where MeHg enters the food chain in
YNP. However, the source of the MeHg in these mats is unknown. However, given that the only
known sources of biological MeHg are through the activity of SRB and IRB populations, it is
possible that one of these organisms may be involved in MeHg production.
The sulfur, iron, and mercury cycles are thought to be intertwined in geothermal
environments. In acid geothermal fluids of volcanic origin solid, aqueous and gaseous forms of
Sulfur Reduction Sulfide Oxidation
Mineralization
Sulfur Oxidation
Figure 3.3. Important
abiotic and
bioticReduction
reactions in the sulfur cycle.
Assimilatory
Sulfate
62
sulfur compounds are commonly found. Solid forms of sulfur are typically formed from abiotic
or biotic H2S oxidation (Figure 3. 3). Abiotic H2S oxidation occurs by reactions with metals such
as vanadate, ferric iron and cupric copper [48]. Abiotic H2S oxidation can also occur when
thiosulfate (S2O32-) is formed and because of the acidic conditions, disproportionates to form
sulfite (SO3-) and elemental sulfur [49]. Biotic H2S oxidation occurs through the actions of sulfide
oxidizing organisms such as Aquificales [50] and proceeds via the thiosulfate and possible sulfite
and elemental sulfur intermediates as described above for the abiotic oxidation of sulfide. For
these reasons, sulfate ions are typically abundant in acidic environments and my serve as an
electron acceptor supporting the activity of Hg methylating SRB. Likewise, Fe is abundant in a
number of geothermal environments in Yellowstone, most notably acidic ecosystems [51] and
thus may serve as an electron acceptor supporting the activity of Hg methylating IRB.
Iron is one of the most common metals found on Earth and is an important metal in
biology for many reasons, especially because of the abundance of proteins containing iron-sulfur
clusters. Iron is found in two oxidation states: ferrous iron, Fe2+(FeII) and ferric iron, Fe3+(FeIII)
[52]. Iron is present as both FeII and FeIII in minerals on Earth’s surface, as FeIII particles in the
atmosphere, and is also found in ionic forms dissolved in water. Iron compounds are produced
by volcanic, anthropogenic and biological activities. FeII minerals are stable at or above neutral
pH in anoxic environments and in aqueous environments FeII is rapidly oxidized by atmospheric
oxygen to form FeIII [52]. FeIII minerals are relatively insoluble at neutral pH values, but are
soluble in acidic environments. In addition elemental iron is oxidized quickly when exposed to
oxygen to form FeII [52].
In YNP the high temperatures and acidic pH of the geothermal systems directly impact
the sulfur and iron cycles because the speciation and stability of a number of sulfur and iron
63
compounds are pH dependent [52, 53]. For example, at low pH iron is stable in its ferrous state
and only through biotic oxidation can FeIII be produced. Likewise, at low pH (<5) the
undissociated form of H2S is the predominant form of sulfide present. H2S is a small, uncharged
molecule, and as such can permeate membranes easily. Within cells, a higher pH is maintained
in which H2S can dissociate, allowing the free sulfide ion to interact with metal ions and
functional groups of electron carrier systems to interfere with cellular metabolism [54]. Because
of these interactions within the cell, H2S is the most toxic form of sulfide [55, 56]. Due to the
challenges that high H2S levels pose on growth in acidic environments, sulfate and sulfur
respiring microorganisms have been the subject of intense study [34, 39, 57, 58].
The geothermal feature in YNP where sampling for this study was completed is referred
to as Dragon spring (unofficial name), an acid-sulfate-chloride (ASC) spring (Figure 3.4). ACS
features are named because of their low pH and high sulfate and chloride concentrations.
Figure 3.4. Dragon Spring sampling site with arrow indicating an
area of Zygogonium mat growth.
64
Dragon Spring is part of the Norris Geyser Basin in YNP, Wyoming, USA (Latitude: 44o
43’54.8’’N, Longitude: 110o 42’39.9’’W) (Figure 3.5). The source water of Dragon Spring has
been measured to have a pH of 3.1 and ranges in temperature from about 66 -800C [59-61].
Many different inorganic energy sources exist in the source water of Dragon Spring, including
H2S, FeII and AsIII as well as dissolved organic carbon [62]. Available electron acceptors include
FeIII after biological oxidation of FeII commences at about 65OC [63], NO3, O2, SO and SO4. While
several studies have detected utilization of most of these redox couples by the Dragon Spring
microbial community, evidence for the utilization of sulfate is lacking, despite numerous efforts
[57]. Moreover, no evidence for FeIII reduction has been demonstrated in these environments.
While evidence for SO4 and FeIII reduction have not been demonstrated in Dragon Spring,
previous studies suggest that the occurrence of these processes is unlikely to be constrained by
the pH of the Dragon Spring environment. Iron reducing bacteria have been found to reduce
soluble ferric iron (FeIII) as well as insoluble ferric oxides [64-66]. At low pH FeIII is more soluble;
studies showing FeIII reduction at low pH are widespread [64, 67]. Likewise, SRB activity has
been identified in many acidic and thermal environments [30, 54, 68, 69]. The purpose of this
study was to identify microbiological populations present in moist sediments beneath
Zygogonium mat communities at Dragon Spring that may participate in the Hg biogeochemical
cycle and which may represent a source of MeHg impacting the geothermal ecosystem. We
hypothesized that either a sulfate- or iron-reducing organism in the sediment beneath the
Zygogonium mats was contributing MeHg to the ecosystem.
65
Norris Geyser Basin
Figure 3.5. Map of Yellowstone showing Norris
Geyser Basin (location of Dragon Spring) sampling
site. Modified from [2].
We employed an enrichment and isolation approach to better understand the potential
for these two functional guilds to contribute to Hg biogeochemical cycling in this acidic
geothermal environment. Here we report the enrichment and partial characterization of a novel
sulfate reducing bacterium from sediment sampled from beneath a Zygogonium mat in the
Norris Geyser Basin of YNP, Wyoming, USA. The results suggest that this population is capable of
utilizing nutrients present in the Dragon Spring environment, including H2, CO2 and SO4. Ongoing
efforts to further purify the novel sulfate reducing bacterium for future physiological,
biochemical, and Hg transformation assays (in collaboration with Professor Tamar Barkay at
Rutgers University) will provide a useful framework for understanding the role of sulfate
reduction in this unique geothermal environment and may provide insight into the interplay of
sulfur and Hg in this environment.
66
Materials and Methods
Sample Collection and Enrichment Conditions
Sediments sampled aseptically from beneath a dense Zygogonium spp. mat inhabiting
Dragon Spring (pH2.6, 30.0OC), Norris Geyser Basin, Yellowstone National Park, Wyoming (44o
43’54.8’’N, 110o 42’39.9’’W) were added to pre-sterilized 160mL serum bottles and were
overlaid with 50 mL of adjacent spring water. The serum bottles were sealed with butyl rubber
stoppers (Geo-Microbial Technologies, Inc., Ochelata, OK) and the bottles and the headspace of
the bottles (~95mL) were purged onsite with N2 for at least 5 min. Killed controls were prepared
by amendment with HgCl2 to a final concentration of 500 μM. Enrichments for iron reducing and
sulfate reducing organisms were made.
Enrichments for iron reducing organisms were made using the following media (per L):
200mL base salts (modified from Sigma Aldrich, St. Louis, MO), 1mM MgSO4, 5μL 1M CaCl2,
100μL Wolfe’s vitamins [70], 300μL SL-10 Trace Metals [71], 125g/L Fe(III) citrate (pH3.0). The
salt solution had the following components (per L): 2.5g NaCl, 5g NH4Cl, 2.5g KH2PO4, 82.3g
citrate pH 3. The salt solution was prepared and autoclaved and the additional components of
the media were filter sterilized and added after autoclaving. Carbon sources including formate,
lactate, citrate and acetate were added to separate, pre-sterilized serum bottles sealed with
sterilized butyl rubber stoppers to a final concentration of 10mM. Samples were inoculated with
500 μL of sediment slurry sample from YNP. Growth of iron reducing organisms was monitored
using a ferrozine assay [72].
Enrichments for sulfate reducing organisms were made using the following medium:
(NH4)2SO4 (0.959g/L), KCl (0.33g/L), CaCl2·2H2O (0.33g/L), MgSO4·7H2O (0.33g/L), KH2PO4
(0.33g/L), yeast extract (0.05g/L), Wolfe’s vitamins (1mL/L) [70] and SL-10 trace metals (1mL/L)
67
[71]; the final concentration of sulfate in the media is 10mM. After addition of all of the salts
and yeast extract, the pH of the growth medium was adjusted to 3.0. Following autoclave
sterilization, filter-sterilized Wolfe’s vitamins and SL-10 trace metals were added [70, 71] as well
as several different carbon sources to a final concentration of 10mM. The carbon sources tested
were acetate, formate, lactate, propionate and H2/CO2 (20%/80% v/v of headspace). The
enrichment bottles were then inoculated with 0.5mL of the Dragon Spring sediment slurry
samples. Hydrogen sulfide was detected using the methylene blue method [70]. Following
detection of hydrogen sulfide in the samples containing H2/CO2, subsequent dilution to
extinction cultures were generated by adding 50 mL of enrichment medium to pre-sterilized
125mL serum bottles which were sealed with pre-sterilized butyl rubber stoppers and purged
with CO2 for 60 min. Following purging, 20% v/v of the headspace was replaced with 100% H2.
Dilution to extinction enrichment cultures were inoculated with 0.5mL of the previous dilution
culture which tested positive for hydrogen sulfide. Multiple iterations of dilution to extinction
resulted in a semi-pure culture, as indicated by epifluorescence microscopy and genetic
analyses. In order to test for the inhibition of sulfate reducing bacteria, at the 5th transfer of the
dilution series, sodium molybdate was added to three serum vials (200μM) containing
enrichment medium and the vials were inoculated with 0.5mL of the previous sulfate reducing
dilution culture. The molybdate cultures were made in parallel with dilution cultures lacking
molybdate.
Epifluorescence Microscopy
To visualize cells using microscopy 1mL of culture was filtered through a 0.02μM, 25mM
filter (Whatman, Mobile, AL) and stained using DAPI blue-fluorescent dye (Life Technologies,
Grand Island, NY) which associates with dsDNA. Cells were visualized using a Nikon Eclipse E600
68
fluorescence microscope (Nikon, Melville, NY) equipped with a Nikon Y-FL epifluorescence unit,
a Nikon plan fluor 40X oil objective, a high pressure Hg vapor lamp and a Nikon UV2A filter
(Nikon). Images were taken using an Olympus Camedia C-4000 4.0MP digital camera with 3X
optical zoom (Olympus Imagine America Inc., Center Valley, PA).
PCR Analysis
PCR amplification of the 16S rDNA gene sequence was performed using a hot start PCR
technique to amplify 16S rDNA gene fragments recovered from enrichment cultures using
bacterial 1100F (5'-YAACGAGCGCAACCC -3') and 1492R (5'- GGTTACCTTGTTACGACTT-3') primers
(Integrated DNA Technologies, Coralville, IA). Template for PCR was obtained by centrifuging
2mL of the enrichment culture and resuspending the pellet in 10μL nuclease free water (Sigma,
St. Louis, MO). Two PCR reaction mixtures were made: pre- and post-hot start solutions. Each
pre-hot start solution contained the following reagents: 3μL resuspended whole cells
(template), 1X PCR buffer (Invitrogen, Grand Island, NY), and nuclease free water (Sigma, St.
Louis, MO). Each post-hot start solution contained the following reactants: 1X PCR buffer
(Invitrogen, Grand Island, NY), nuclease free water (Sigma, St. Louis, MO), 2 mM MgCl2
(Invitrogen, Grand Island, NY), 200 µM of each deoxynucleotide triphosphate (Eppendorf,
Hamburg, Germany), 0.5 µM of each forward and reverse primer, 0.4 µg µL-1 molecular-grade
bovine serum albumin (Roche, Indianapolis, IN) and 0.25 U Taq DNA Polymerase (Invitrogen,
Grand Island, NY) to a final volume of 50 µL. PCR was performed according to the following
conditions: Pre-hot start solution (99oC, 15 min.), followed by addition of post-hot start solution
to reach a total volume of 50 μL. After addition of post-hot start solution the following
conditions were used: initial denaturation (94°C, 4 min.) followed by 35 cycles of: denaturation
69
(94°C, 1 min.), annealing (55°C, 1 min.), and elongation (72°C, 1.5 min.) with a final extension
step at 72°C for 20 min.
16S rDNA Gene Analysis
PCR amplicons were purified using the Wizard PCR Preps DNA Purification System
(Promega, Madison, WI) cloned, assembled, and analyzed according to previously published
protocols [73] using the pGEM-T Easy Vector (Promega, Madison, WI). The phylogenetic
affiliation of sequences was determined using BLASTn.
Results and Discussion
Dragon Spring Site Description
The solid and aqueous phase chemistry of Dragon Spring has been reported previously
[62]. On the day of sampling (August, 2011) the pH and temperature of the source water was
measured as 2.86 and 78oC, respectively. Sampling was completed approximately 10m from the
source of the spring where the temperature was about 26OC and the pH was 2.6. At the
sampling site there was a large Zygogonium mat (Figure 3.4).
Dragon Spring Isolate Enrichment and Isolation
Media for the selective enrichment of iron reducing microorganisms inoculated with
Dragon Spring sediment slurry sample and stored at 20oC tested negative for the presence of FeII
following several weeks of incubation, indicating an iron reducing microorganism was most
likely not present or active under the conditions in the enrichment medium. Medium containing
H2/CO2 as the electron and carbon sources for the selective enrichment of sulfate reducing
microorganisms inoculated with Dragon Spring sediment slurry sample and stored at 20oC
70
tested positive for the presence of H2S after 2 weeks, indicating a sulfate reducing
microorganism was present and active in the culture. Hydrogen sulfide was detected in the
aqueous phase of biological replicates, while no activity was observed in the killed controls,
suggesting that the sulfate reducing activity was attributable to microbial activity and not abiotic
reduction [62]. To make a pure culture, iterations of dilution to extinction were completed on
the culture which tested positive for H2S. In order to test for inhibition of a sulfate reducing
microorganism, samples containing sodium molybdate were made in parallel with samples
lacking molybdate following the 5th dilution to extinction. Hydrogen sulfide was never detected
in the samples containing molybdate, while it was detected in the live samples. These results
indicate a sulfate reducing organism is present, and inhibited by the presence of molybdate. The
evidence that sulfate reduction is inhibited by molybdate in these samples means that in future
tests of the ability of the isolated microorganisms to methylate Hg, sulfate reduction can be
inhibited easily to see if Hg methylation still occurs. These future studies will indicate if Hg
methylation is due to the activity of a sulfate reducing microorganism or is abiotic (which would
be shown if Hg methylation occurs in the presence of molybdate).
Negative control
no template
Positive control
bacterial 16S
Dragon Spring
Isolate 16S
Following 8 rounds of dilution to extinction, 16S rDNA genes were amplified using a
1Kb
Ladder
500bp
Figure 3.6. PCR amplicons of bacterial 16S genes.
71
universal primer set for bacterial 16S rDNA, sequenced and characterized by BLASTn analysis.
PCR amplification using a universal primer set for archaeal 16S rDNA yielded no products,
evidence supporting the presence of a SRB. Analysis of PCR-amplified 16S rDNA genes indicated
several 16S rDNA gene phylotypes from the final dilution culture (Figure 3.6). 16S rDNA gene
fragments (corresponding to positions 1100-1492 of the E. coli 16S rDNA gene) were used for
genetic analysis of the Dragon Spring culture.
The sequencing results indicated the presence of three different bacterial species:
Paludibacter propionicigenes WB4 strain WB4, Thiomonas intermedia K12, and
Thermodesulfobium narugense DSM 14796. The percent identity that each of these species has
with the 16S sequences from the environmental cultures is 92%, 100% and 99%, respectively.
Each of these species has been shown to reduce sulfate during growth. Paludibacter
propionicigenes WB4 strain WB4 was first isolated in 2006 from plant residue in a Japan rice
field [74]. The organism was found to grow anaerobically at an optimal temperature of 30OC and
a pH of 6.6 [74]. Based on the growth conditions of the Dragon Spring enrichment culture and
that of Paludibacter propionicigenes WB4 strain WB4, it is possible that a species of Paludibacter
is present in the enrichment and actively reducing sulfate, although the other bacterial species
found to be present in the enrichment cultures have optimal growth conditions more similar to
the enrichment conditions. Thiomonas intermidia K12 was first isolated from a sewer wall in
Hamburg, Germany [75]. However, this isolate is optimally grown at pH values between 4.7 and
6.6 and at a temperature of 30OC; these conditions are not similar to the conditions used in the
Dragon Spring enrichment, meaning one of the Dragon Spring isolates is most likely not
Thiomonas intermidia K12, but could be related [75, 76]. Thermodesulfobium narugense DSM
14796 was first isolated from a hot spring in Narugo, Japan [69] and found to grow anaerobically
72
using hydrogen and carbon dioxide by sulfate reduction. T. narugense was found to grow
optimally at pH 4.0-6.5 and a temperature of 50-55OC [69]. Based on the similarity between the
growth conditions of the Dragon Spring enrichment culture and those of Thermodesulfobium
narugense, it is possible that Thermodesulfobium narugense is the organism responsible for
reducing sulfate in the enrichment culture.
Pairwise alignment of the three 16S rDNA sequences from the Dragon Springs
enrichment sample yielded similarities of 51%, 34% and 29%, suggesting the three sequences
were from unrelated bacterial species. The sequence data supports the presence of a semi-pure
culture that, based on previous research, may contain bacterial species related to Paludibacter
propionicigenes WB4 strain WB4, Thiomonas intermedia K12, and Thermodesulfobium
narugense DSM 14796 [69, 75, 77]. Continued work on this project should involve further
iterations of dilution to extinction of the current enrichment cultures, with the goal of isolating a
single sulfate reducing organism.
Morphology of Dragon Spring Isolate
Epifluorescence microscopy and imaging revealed rod shaped cells, all of similar
morphology (Figure 3.7). These imaging results contrast with what is seen by genetic analysis of
16S rDNA samples which suggest the presence of more than one species of bacteria that are
genetically dissimilar.
73
Figure 3.7. Epifluorescent image of Dragon Spring enrichment
culture using Dapi stain.
In summary, the results presented here indicate the presence of a sulfate reducing
organism isolated from beneath a Zygogonium mat at the Dragon Spring hot spring in
Yellowstone National Park. Further iterations of dilution to extinction experiments, which are
ongoing, need to be completed to generate an axenic culture which can then be characterized
through growth and biochemical studies and which then can be tested for Hg methylation
activity. The finding that molybdate inhibits sulfate reduction in the enrichment culture is
important because it will allow for future studies of mercury methylation at the mat. For
example, a future study would be to make primers specific for the bacterial species that is finally
isolated from the current enrichment samples. Next, sediment slurry samples would be made as
before, but with sulfate added to all samples and molybdate or glutaraldehyde to controls. The
presence of sulfate would allow any sulfate reducing organisms to be active. The molybdate
would serve as a control to knock out biological sulfate reduction and rule out abiotic sulfate
reduction. Finally, the glutaraldehyde samples would provide killed controls which also tests for
74
the presence of abiotic sulfate reduction. Mercury methylation activity could then be assessed
in each sample, as well as genetic analyses. Genetic analyses would use primers specific for the
isolate from the current dilution to extinction enrichment samples. PCR of the sulfate,
molybdate and glutaradehyde containing cultures using these primers and subsequent
sequencing would indicate the culture currently being isolated contains the bacterium
responsible for Hg methylation in the Zygogonium mats. Subsequent genetic analyses (e.g.,
qPCR and qRT-PCR) can then be used to better understand the temporal and spatial dynamics of
this anaerobic population(s) in the sediments beneath an oxygenic, phototrophic mat and the
role that this population(s) may have in generating MeHg under these conditions.
75
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82
CHAPTER 4
CONCLUSIONS
In conclusion, the work presented herein provides insights into microbial metabolism
through the work completed on three separate projects involving the nitrogenase homologs
from Roseiflexus castenholzii, novel enzymes involved in the tricarboxylic acid (TCA) cycle in
cyanobacteria, and enrichment of sulfate reducing organisms from a hot spring in Yellowstone
National Park (YNP). Results from growth experiments of R. castenholzii in which fixed nitrogen
was not included in the media suggest that R. castenholzii is not able to fix nitrogen in the
presence of the carbon sources tested (acetate, citrate, glucose, glycyl-glycine, pyruvate).
Additional carbon sources should be tested and expression studies using qRT-PCR should be
used in order to confirm that R. castenholzii cannot fix nitrogen and determine if the level of
expression of the nitrogenase homolog genes is changing during growth without fixed nitrogen.
The growth studies provide initial results suggesting that R. castenholzii cannot grow without
fixed nitrogen, however the experiments do not suggest a role for the nitrogenase homologs in
the organism. In order to determine if heterologous expression in E. coli was a tractable system
for overexpression of the nitrogenase homologs, expression experiments were carried out.
Results suggest that NifH and NifDK proteins are overexpressed in E. coli and are able to be
partially purified. However, with all of the proteins that were overexpressed, solubility is an
issue and additional buffer conditions need to be explored in order to keep the proteins soluble
for biochemical assays and crystallization trials. Initial results suggest that the NifB protein is not
being overexpressed in the E. coli host which is a problem that could be solved by changing the
growth conditions (for example: lowering expression temperature, raising or lowering the pH of
83
the media or using an alternate type of media) or changing the construct of the protein in the
expression vector.
Results from the study of three novel TCA enzymes from the cyanobacterial species
Synechococcus sp. PCC 7002 provide working protocols for expression and purification of both
the novel fumarase and succinic semialdehyde dehydrogenase (SSADH). In addition to
expressing and purifying these two proteins, initial crystallization screens were tested, however,
no crystal growth was observed. With working expression and purification protocols, attempts
to crystallize the proteins in alternative buffers will be simplified. The third novel TCA enzyme,
the 2-oxoglutarate decarboxylase (2-OGDC) was not overexpressed in E. coli at levels that were
sufficient for further purification and crystallization screens.
The third set of experiments which probed the source of methylmercury in Zygogonium
mats in YNP resulted in an enrichment culture containing at least three sulfate reducing
bacteria. Sulfate reducing bacteria are one of two guilds of organisms which have been shown
previously to methylate mercury. Further iterations f dilution to extinction need to be
completed in order to enrich for an axenic culture. With a pure culture, many different
experiments can be completed in order to find the conditions for optimal growth of the
bacterium and also to explore the role of the bacterium isolated in mercury methylation in YNP.
In order to test if the isolated bacterium is responsible for mercury methylation in the
Zygogonium mats, samples can be taken from the hot spring and manipulated by adding sulfate,
molybdate (an inhibitor of sulfate reduction), methylmercury, mercury and mercuric chloride.
Then 16S rDNA samples can be taken from the samples and genetic analysis will reveal what
organisms are present; qRT-PCR on the samples using primers specific for the isolated sulfate
reducing bacterium will indicate if the bacterium is active in the samples. These experiments will
84
provide evidence regarding the organism(s) which are responsible for the entry of
methylmercury into Zygogonium mats in YNP. This project, in combination with the previous
two projects, provides important initial results that can be used in continuing studies on the
metabolic processes involved.
85
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