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IDENTIFICATION OF UNKNOWN METHIONINE SULFOXIDE REDUCTASE ACTIVITY IN HALOFERAX VOLCANII
By
ZACHARY ADAMS
A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE
UNIVERSITY OF FLORIDA
2018
© 2018 Zachary Adams
To Mom and Dad
4
ACKNOWLEDGMENTS
First and foremost, I thank my parents for their love and support. I thank my
mentor Dr. Julie Maupin-Furlow, as well as my committee members Dr. Tony Romeo
and Dr. Christopher Reisch for their guidance. I am also grateful to the faculty, staff, and
my colleagues in the Microbiology and Cell Science department.
5
TABLE OF CONTENTS page
ACKNOWLEDGMENTS .................................................................................................. 4
LIST OF TABLES ............................................................................................................ 6
LIST OF FIGURES .......................................................................................................... 7
ABSTRACT ..................................................................................................................... 8
1 INTRODUCTION .................................................................................................... 10
Oxidative Damage .................................................................................................. 10
Methionine Oxidation and Repair ............................................................................ 10 Distribution of Methionine Sulfoxide Reductases .................................................... 11
Molybdopterin Dependent Reduction of Methionine Sulfoxide ................................ 12 Molybdenum Cofactor Biosynthesis ........................................................................ 13
2 PURPOSE .............................................................................................................. 18
3 METHODS .............................................................................................................. 21
Strains and Culture Conditions ............................................................................... 21
General DNA Methodology ..................................................................................... 21
Construction of Deletion Plasmids .......................................................................... 22 Generation of Mutant Strains .................................................................................. 23 Growth Assays ........................................................................................................ 23
4 RESULTS AND DISCUSSION ............................................................................... 29
Deletion of moaE .................................................................................................... 29
ZA106 Fails to Utilize Methionine Sulfoxide ............................................................ 29 Deletion of dmsA .................................................................................................... 30 ZA109 Retains Ability to Grow on Methionine Sulfoxide ......................................... 31 Identification and Deletion of Additional Candidates ............................................... 31
Plate Assay Using Candidate Gene Deletion Strains ............................................. 32
5 CONCLUSIONS ..................................................................................................... 46
LIST OF REFERENCES ............................................................................................... 48
BIOGRAPHICAL SKETCH ............................................................................................ 52
6
LIST OF TABLES
Table page 2-1 List of strains used in this study .......................................................................... 25
2-2 List of plasmids used in this study ...................................................................... 26
2-3 List of primers used in this study ........................................................................ 27
7
LIST OF FIGURES
Figure page 1-1 Repair of methionine sulfoxide by MsrA and MsrB ............................................. 15
1-2 Overview of Moco biosynthesis and its derivatives bis-MGD and MCD in E. coli ...................................................................................................................... 16
1-3 Working model of protein conjugation and sulfur transfer pathways in archaea .............................................................................................................. 17
4-1 PCR screening confirming deletion of moaE in strain ZA106 ............................. 34
4-2 Growth assay comparing strains H26, XF127, XF130, and ZA106 .................... 36
4-3 PCR screening confirming deletion of dmsA in strain ZA109 ............................. 39
4-4 Growth assay comparing strains H26, XF127, XF130, ZA106, and ZA109 ........ 41
4-5 Summary of additional candidate gene information and their respective proteins ............................................................................................................... 43
4-6 PCR screening of candidate gene deletion isolates for strains ZA110-116 ........ 44
4-7 Plate assay for growth of additional candidate mutant strains on MetSO ........... 45
8
Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science
IDENTIFICATION OF UNKNOWN METHIONINE SULFOXIDE REDUCTASE ACTIVITY
IN HALOFERAX VOLCANII
By
Zachary Adams
December 2018
Chair: Julie A. Maupin Major: Microbiology and Cell Science
The major methionine sulfoxide reductase enzymes MsrA and MsrB are well
conserved across all domains of life and seem to have evolved convergently out of the
necessity for life in an oxygen-rich world. The apparent lack of these of enzymes in the
majority of thermophilic archaea, among others, is not fully understood.
Recent observations in the archaeon Haloferax volcanii indicate that a
methionine auxotroph with both predicted methionine sulfoxide reductase genes deleted
is still capable of utilizing methionine sulfoxide for growth. This finding reveals that
additional methionine sulfoxide reductase activity is present in H. volcanii and yet to be
identified. We set out to identify which enzyme(s) were responsible for such activity.
While MsrA/B utilize a nucleophilic active site cysteine for reduction of
methionine sulfoxide (MetSO), some oxidoreductases capable of reducing MetSO utilize
molybdopterin based cofactors. To determine if the yet to be identified Msr activity in H.
volcanii was molybdopterin dependent, we further deleted the moaE gene, proposed to
play a key role in biosynthesis of molybdopterin. Accordingly, the resulting disruption of
molybdopterin biosynthesis was found to abolish growth on methionine sulfoxide. We
then identified 8 putative molybdopterin oxidoreductases in H. volcanii and deleted
9
these genes in the initial strain. Work is ongoing to test these deletion strains for
utilization of methionine sulfoxide and is anticipated to serve in the identification of the
unknown methionine sulfoxide reductase(s) in this organism. Such a discovery would
further our knowledge of this increasingly diverse group of enzymes and their unusual
distribution throughout archaea.
10
CHAPTER 1 INTRODUCTION
Oxidative Damage
While the development of Earth’s oxygen rich atmosphere certainly led to an
explosion of life and molded life as we know it today, oxygen also presents unique
challenges to living organisms. Both endogenous factors, such as metabolism, and
exogenous factors, like radiation, can lead to the formation of free radicals and reactive
oxygen species. The formation of these reactive oxidants and the damage they can
cause in living organisms has been extensively reviewed1. Highly reactive oxidants can
react with essentially any biological molecule, including nucleic acids, proteins, lipids,
and carbohydrates. Most early research on the oxidation of biomolecules focused on
lipids and DNA, but scientific advances have addressed difficulties in studying the
oxidation of proteins. Given that proteins constitute the major biological component of
living systems by weight and are estimated to consume the majority of radicals in cells,
the study of protein oxidation represents a critical field2.
Methionine Oxidation and Repair
The sulfur containing amino acids cysteine and methionine (Met) are readily
oxidized by a variety of reactive oxidants. While the unique thiol chemistry of cysteine
has long been understood to play major roles in redox sensing and regulation, protein
folding and function, and others, the importance of methionine oxidation did not gain
much interest until recently3. Oxidation of methionine forms two diastereomers of
methionine sulfoxide (MetSO) in equal measure, Met-(R)-SO and Met-(S)-SO4. Further
oxidation of MetSO is possible, giving methionine sulfone, but this rarely occurs under
11
typical conditions. Methionine oxidation can occur both on free methionine and residues
in protein3,5.
Repair of MetSO back to methionine is possible using methionine sulfoxide
reductase (Msr) enzymes. The two major types of Msrs are MsrA and MsrB. MsrA type
enzymes specifically reduce the Met-(S)-SO stereoisomer both in the free and protein
bound form. MsrB enzymes reduce Met-(R)-SO, also in the free and protein bound
forms, however activity towards free Met-(R)-SO is often much lower6-9. Intriguingly,
MsrA and MsrB share little to no sequence or structural homology yet possess mirror
image active sites and appear to have developed through convergent evolution10,11.
Another class of Msr is represented by enzymes specific for the reduction of free Met-
(R)-SO, the free methionine-(R)-sulfoxide reductase or fRMsr12,13. An overview of the
mechanism of repair by Msrs is shown in Figure 1-1.
Distribution of Methionine Sulfoxide Reductases
As early as the initial discoveries of MsrA and MsrB, it had become evident that
Msr enzymes were represented in all domains of life14,15. Soon thereafter, the dawn of
the post-genomic era confirmed the widespread distribution of Msrs. All eukaryotes and
cyanobacteria examined to date are predicted to encode for MsrA and MsrB, with
multiple isoforms often present16,17. Most bacteria also possess MsrA and MsrB,
however there are some exceptions, including some established bacterial
endosymbionts and endoparasites16,17. Free-living bacteria in which both Msrs are
predicted missing are restricted to a small number of anaerobes and
hyperthermophiles16-18. It is also worth noting that in many bacteria, MsrA and MsrB
domains are fused together within a single protein16,17,19.
12
Among the three domains of life, Msrs appear to be most scarcely distributed in
the archaea. Many archaea are predicted to lack MsrB, and most thermophiles are
predicted to lack both MsrA and MsrB16-18. A notable exception is the hyperthermophilic
archaeon Thermococcus kodakaraensis, which grows at an optimal temperature of 85
°C20. However, the MsrA/B fusion protein of this organism could not be detected in vivo
at temperatures 80-90 °C, and maximum MsrA/B activity was observed at 30 °C18.
Fukushima et. al. proposed that lower oxygen solubility in high temperature
environments may eliminate the need for MetSO reduction and that T. kodakaraensis
may benefit from this enzyme in lower temperature environments18. This hypothesis
may be reasonable given the temperature gradients located at hydrothermal vents but
does not explain if or how other archaea which lack predicted Msrs might cope with
increasing dissolved oxygen at lower temperatures.
Molybdopterin Dependent Reduction of Methionine Sulfoxide
Methionine sulfoxide can also be reduced by oxidoreductases that act on N- and
S-oxide substrates with broad specificity. One example is DMSO reductase, which has
been shown to exhibit broad substrate specificity in bacteria21. In some instances,
molybdopterin dependent enzymes of the DMSO reductase family may display
considerably high activity towards MetSO. E. coli BisC is one such enzyme, in which the
first enzymatic activity for the stereospecific reduction of free MetSO was observed,
allowing E. coli to use free Met-(S)-O for growth22. More recently, MsrP, a novel Msr
enzyme capable of reducing all forms of MetSO with the help of its partner MsrQ, was
identified in the bacterial cell envelope23. Although a member of the molybdopterin
dependent sulfite-oxidase family, homologs of MsrP appear to be strictly contained in
bacteria24. The common feature amongst these enzymes is that they use variants of
13
molybdenum cofactor for reduction of MetSO, as opposed to active site cysteines found
in the classical MsrA, MsrB, and fRMsr9,24. Overall, molybdopterin-dependent
oxidoreductases appear to be playing a larger and larger role in our still growing
understanding of MetSO reduction.
Molybdenum Cofactor Biosynthesis
Biosynthesis of the molybdenum cofactor (Moco) is carried out in three steps.
The first step involves cyclization of GTP to form cyclopyranopterin monophosphate
(cPMP). cPMP is then sulfurylated resulting in the mature form of pyranopterin (MPT).
This step is carried out by MPT synthase, a heterotetrameric complex of MoaD and
MoaE. Following adenylation of MPT, a molybdate ion is inserted to give the most basic
form of Moco (or more informatively, Mo-MPT) catalyzed by the MogA and MoeA
proteins25-27.
Moco can be modified to give derived forms of the cofactor, such as through
addition of GMP or cytosine. Two molecules of the guanine modified Moco (MGD or
MPT guanine dinucleotide cofactor) are joined around one molybdenum center to form
bis-MGD28. bis-MGD is found in enzymes of the DMSO reductase family in bacteria,
whereas a cytosine modified form (MCD) is found in enzymes of the xanthine oxidase
family in E. coli27. While MobA is essential for the attachment of GMP to Moco29, the
function of MobB remains to be determined. Initial reports suggested that MobB might
function as an adapter protein that assists MobA through binding of GTP30. However, a
more recent study showed in vitro that MobA alone is sufficient for formation and
insertion of bis-MGD in the DMSO reductase of Rhodobacter sphaeroides31. See Figure
1-2 for details.
14
In Haloferax volcanii, the MoaE homologue has been observed in complex with
small archaeal modifier protein 1 (SAMP1)32. Given the similarity of H. volcanii MoaE
and SAMP1 to the MPT synthase subunits of other organisms, MoaE is proposed to
function in the sulfurylation of cPMP to form molybdopterin, using thiocarboxylated
SAMP1 as a source of sulfur. This proposal is further supported by the finding that
MoaE is essential for DMSO reductase activity in H. volcanii33. Furthermore, MoaE is
found fused to an N-terminal MobB domain, and this arrangement is often observed in
other halophilic and methanogenic archaea33. A working model for the protein
conjugation and sulfur transfer pathways of archaea is shown in Figure 1-3.
15
Figure 1-1. Repair of methionine sulfoxide by MsrA and MsrB. Reprinted by permission
from Springer Nature: Nature Reviews Microbiology, Oxidative stress, protein damage and repair in bacteria, Gennaris, A., Barras, F. & Collet, J.-F9, Copyright © 2017.
16
Figure 1-2. Overview of Moco biosynthesis and its derivatives bis-MGD and MCD in E.
coli. Reprinted from Biochimica et Biophysica Acta (BBA) – Bioenergetics, 1827, Iobbi-Nivol, C. & Leimkühler, S., Molybdenum enzymes, their maturation and molybdenum cofactor biosynthesis in Escherichia coli, 1086–110127, Copyright © 2013, with permission from Elsevier.
17
Figure 1-3. Working model of protein conjugation and sulfur transfer pathways in
archaea. Figure courtesy of: Miranda, H. V. et al. E1- and ubiquitin-like proteins provide a direct link between protein conjugation and sulfur transfer in archaea. Proceedings of the National Academy of Sciences 108, 4417–4422 (2011)33.
18
CHAPTER 2 PURPOSE
The absence of a predicted Msr in many archaeal species frames the question of
how these organisms seemingly cope with an inability to reduce MetSO in free or
protein form. Only a handful of Msrs from the archaea have been characterized24. This
group of studied archaeal Msrs includes the previously mentioned MsrAB fusion protein
from T. kodakaraensis18. In addition, the MsrB of Methanothermobacter
thermoautotrophicus10, and an fRMsr from Thermoplasma acidophilum34, are
biochemically and structurally characterized. Lastly, the MsrA and MsrB of Haloferax
volcanii were found to reduce the peptide mimic dabsyl-Met-(S)-O and -Met-(R)-O,
respectively35. Interestingly, treatment of H. volcanii with DMSO stimulates a MsrA-
dependent ubiquitin-like conjugation process, accompanied by an inhibition of the
MetSO reductase activity of MsrA35.
Haloferax volcanii is a halophilic archaeon belonging to the phylum
Euryarchaeota. Originally isolated from the Dead Sea, it grows optimally in NaCl
concentrations from 1.7-2.5 M and at temperatures near 42 °C36. Compared to other
archaea, these relatively moderate growth conditions of H. volcanii make it readily
suitable for laboratory study. Furthermore, the development of a variety of genetic,
biochemical, and now “omics” based tools have led to this organism’s standing as one
of the foremost model organisms in the study of archaea37,38. In light of recent findings
regarding MsrA of H. volcanii, this system represents a promising subject for the study
of Msrs in archaea.
MsrA and MsrB are the only predicted Msrs present in H. volcanii. Our efforts, to
confirm MsrA and MsrB were the only Msrs present in this organism, led to the
19
surprising observation that a methionine auxotroph lacking both MsrA and MsrB can still
utilize MetSO as its only source of methionine for growth. This finding suggests that
additional Msr activity is present in H. volcanii. Given that no dabsyl-MetSO activity was
detected in an MsrAB mutant35, it appears that residual Msr activity in H. volcanii is
likely to be specific for free MetSO.
Although fRMsrs are conserved to a degree in gram-negative bacteria, they
appear to be absent in the vast majority of archaea and are not predicted in any
Halobacteria genomes24. Investigation of this residual Msr activity in H. volcanii may
shed light on whether specific reduction of free MetSO occurs in diverse archaea. In
addition, knowledge of the enzyme(s) involved may provide clues as to other Msrs yet
to identified, perhaps in thermophilic archaea, for instance. Furthermore, this knowledge
would help bolster our understanding of Msrs in a leading model organism for archaea,
where it has already been shown that MsrA plays a complex role in response to
DMSO35.
Our primary objective of this study is to identify the enzyme(s) responsible for the
residual Msr activity identified in H. volcanii. As an initial screen to determine whether
residual activity is molybdopterin dependent, we disrupted the synthesis of Moco
through deletion of moaE from the H26 derived parent strain XF130 (ΔmetE1/2 ΔmsrA
ΔmsrB). Utilization of MetSO was abolished in this strain. This provided evidence that
any residual Msr activity—at a level sufficient to support growth on MetSO—was
molybdopterin dependent. The discovery that E. coli BisC carries out the stereospecific
reduction of free Met-(S)-O was made using a similar approach22. As this approach was
successful, we now use bioinformatic analysis to predict all molybdopterin
20
oxidoreductases in the genome, and candidates were chosen for deletion from the
parent strain. Selection of the enzyme responsible for residual Msr activity would result
in a failure to utilize MetSO for growth. Early follow up experiments will focus on
characterization of the enzyme and its activity towards reduction of MetSO.
21
CHAPTER 3 METHODS
Strains and Culture Conditions
Strains used in this study are listed in Table 2-1. E. coli TOP10 was used for
routine selection, amplification, and maintenance of plasmid DNA. E. coli GM2163 was
used for isolation of plasmid DNA prior to transformation of H. volcanii. E. coli strains
were grown in LB-Miller medium at 37°C. H. volcanii strains were grown at 42-45°C in
ATCC 974, minimal medium (Hv-Min), glycerol minimal medium (GMM), and casamino
acids medium (HvCa) as previously described39,40. Solid medium was supplemented
with 1.5 and 2.0 % (wt/vol) agar for culture of E. coli and H. volcanii, respectively.
Ampicillin was added to LB medium at a concentration of 100 ug/mL where necessary.
Liquid cultures were aerated by rotary shaking at 200 rpm. Cells were stored at -80°C in
20% (vol/vol) glycerol stocks. H. volcanii strains were streaked from the freezer stocks
onto ATCC 974 agar plates.
General DNA Methodology
Plasmids used in this study are listed in Table 2-2. Primers are listed in Table 2-
3. Phusion DNA polymerase was used for high-fidelity amplification of DNA, while
OneTaq and/or Phusion DNA polymerase was used for screening purposes, according
to the manufacturer’s instructions (New England Biolabs). PCR products were analyzed
on 0.8 % (w/v) agarose gels stained with 0.5 μg/mL ethidium bromide in Tris-acetate-
EDTA (TAE) buffer. For PCR products intended for downstream gel extraction and
restriction digest, agarose gels were stained with GelGreen according to the supplier’s
instructions (Biotium). Restriction enzymes, ligase mix, and KLD (kinase, ligase, DpnI)
mix were used according to the manufacturer’s instructions (New England Biolabs).
22
DNA cleanup, gel extraction, and plasmid minipreps were completed using the
respective Monarch purification kits (New England Biolabs). Sanger sequencing was
used to verify the integrity of all constructs (Eton Biosciences).
Construction of Deletion Plasmids
Primers flanking the gene of interest by roughly 500 base pairs on both sides
were designed and checked for specificity using Primer-BLAST (NCBI)41. Restriction
sites and additional bases for optimal cutting were selected for compatibility with
plasmid pTA131 and added onto the primers found specific by Primer-BLAST. These
primes were named “gene/HVO_xxxx pKO UP/DN” (where HVO_xxxx represents the
locus tag number for each respective gene). Inserts for pre-deletion plasmids were
obtained by PCR amplification using the 500 base pair flanking primers, followed by gel
extraction and restriction digest with the appropriate enzyme. The cleaved inserts were
ligated with cut and phosphatase treated pTA131 to form the pre-deletion plasmid. Pre-
deletion plasmids were transformed to E. coli TOP10, screened, and sequenced.
Inverse PCR primers were designed amplifying outwards from the gene of
interest in the pre-deletion plasmid using Primer3Plus42, and named “gene/HVO_xxxx
UP/DN INV.” Where possible, primers were designed for clean deletion of the gene from
the chromosome, or to at least have minimal impact on adjacent genetic material. The
linear product of the inverse PCR reaction amplifying the pre-deletion plasmid was
treated with KLD (kinase, ligase, DpnI) enzyme mix. The resulting deletion plasmid was
transformed to E. coli TOP10. After verification of the construct by DNA sequencing,
deletion plasmids were passaged through E. coli GM2163 before transformation to H.
volcanii strains.
23
Generation of Mutant Strains
H. volcanii mutants containing chromosomal deletions were generated using the
pyrE2-based “pop-in pop-out” method described previously43,44. Deletion plasmid
integrants were selected on HvCa+ agar without uracil. Screening was conducted using
the “pKO UP/DN” primers, and isolates with PCR products indicative of integration were
selected for the “pop-out” step. Isolates were inoculated in 3 mL ATCC 974
supplemented with 5-FOA and cultured for 24 h. Cells (1.8 mL) were harvested by
centrifugation at 6,000 x g for 10 min at room temperature and resuspended in 0.5 mL
18% saline water (SW)39. Serial dilutions (100, 10-1, 10-2 ) were made, and 50 μL of
each dilution was plated on HvCa agar supplemented with 10 μg/mL uracil and 50
μg/mL 5-FOA. Plates were incubated for 5 days at 42 °C, and colonies were patched on
the same medium for screening. Initial screening was conducted using the 500 base
pair flanking primers, and isolates indicating loss of the gene were chosen. Selected
pop-outs were then streaked for further isolation and gene deletion was confirmed by
screening with a different flanking primer pair and/or gene specific primers.
Growth Assays
Strains selected for growth assays in 96-well plates (CellPro, Alkali Scientific
catalog #TPN1096-NT) were inoculated in 2.5 mL GMM supplemented with 0.1 mM L-
methionine and grown to log phase (OD600 of 0.6) in borosilicate glass 13 x 100 mm test
tubes. The log phase cells were subcultured to an initial OD600 of 0.06 in 2.5 mL of the
same medium and grown to OD600 of 0.6. Cells (0.5 mL of this culture) were pelleted by
centrifugation at 6,000 x g for 10 min at room temperature, and 0.48 mL of the
supernatant was carefully removed. A volume of GMM was added so that 5 μL of the
suspension contained 0.003 OD600 units cells. Aliquots (5 μL) of this suspension were
24
used to inoculate each well of a 96-well plate containing 145 μL GMM supplemented
with varying amounts of L-Met or L-MetSO (Sigma-Aldrich, catalog #M1126). Strains
were inoculated in triplicate and positioned to minimize effects of evaporation across
strains and replicates. The outermost wells were filled with 250 μL water to limit
evaporation of the inner wells. Plates were covered with lids and incubated in a Synergy
HTX plate reader (BioTek) with maximum orbital pattern shaking at 42 °C. OD readings
were taken at 600 nm every one or two hours. Roughly every 24 h, the read was
paused, and the plate was removed briefly for replenishing of the water in the outermost
wells. In the second growth assay containing strain ZA109, water was not replaced in
the outermost wells.
For the growth assay on solid medium, strains were picked from the selective
medium used for pop-out described above and inoculated directly on Hv-Min agar
supplemented with 0.25 mM Met or MetSO. When supplemented with Met, the plate
was incubated for 42 h, whereas with MetSO supplementation, incubation lasted a
period of 9 days.
25
Table 2-1. List of strains used in this study.
Strain Description Source or reference
E. coli
TOP10 F- recA1 endA1 hsdR17(rK– mK+) supE44 thi-1 gyrA relA1
Invitrogen
GM2163 F- ara-14 leuB6 fhuA31 lacY1 tsx78 glnV44 galK2 galT22 mcrA dcm-6 hisG4 rfbD1 rpsL136 dam13::Tn9 xylA5 mtl-1 thi-1 mcrB1 hsdR2
New England Biolabs
H. volcanii
DS70 wild type isolate DS2 cured of plasmid pHV2 Wendoloski et. al., 2001
H26 DS70 ΔpyrE2 Allers et. al., 2004
XF127 H26 ΔmetE1/2 This study
XF128 H26 ΔmetE1/2 ΔmsrA This study
XF129 H26 ΔmetE1/2 ΔmsrB This study
XF130 H26 ΔmetE1/2 ΔmsrA ΔmsrB This study
ZA106 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔmoaE This study
ZA109 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔdmsA This study
ZA110 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔHVO_0671 This study
ZA111 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔHVO_1471 This study
ZA112 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔHVO_B0367 This study
ZA113 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔHVO_B0164 This study
ZA114 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔHVO_0935 This study
ZA115 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔHVO_1908 This study
ZA116 H26 ΔmetE1/2 ΔmsrA ΔmsrB ΔHVO_B0235 This study
26
Table 2-2. List of plasmids used in this study
Plasmid Description Source or reference
pTA131 Apr; Nvr; pyrE2-based integration vector Allers et. al., 2004
pJAM1114 Apr; Nvr; pTA131-based deletion vector for moaE Miranda et. al., 2011
pJAM3500 Apr; Nvr; pTA131-based pre-deletion vector for dmsA
This study
pJAM3501 Apr; Nvr; pTA131-based deletion vector for dmsA This study
pJAM3502 Apr; Nvr; pTA131-based pre-deletion vector for HVO_0671
This study
pJAM3503 Apr; Nvr; pTA131-based deletion vector for HVO_0671
This study
pJAM3504 Apr; Nvr; pTA131-based pre-deletion vector for HVO_1471
This study
pJAM3505 Apr; Nvr; pTA131-based deletion vector for HVO_1471
This study
pJAM3506 Apr; Nvr; pTA131-based pre-deletion vector for HVO_B0367
This study
pJAM3507 Apr; Nvr; pTA131-based deletion vector for HVO_B0367
This study
pJAM3508 Apr; Nvr; pTA131-based pre-deletion vector for HVO_B0164
This study
pJAM3509 Apr; Nvr; pTA131-based deletion vector for HVO_B0164
This study
pJAM3510 Apr; Nvr; pTA131-based pre-deletion vector for HVO_0935
This study
pJAM3511 Apr; Nvr; pTA131-based deletion vector for HVO_0935
This study
pJAM3512 Apr; Nvr; pTA131-based pre-deletion vector for HVO_1908
This study
pJAM3513 Apr; Nvr; pTA131-based deletion vector for HVO_1908
This study
pJAM3514 Apr; Nvr; pTA131-based pre-deletion vector for HVO_B0235
This study
pJAM3515 Apr; Nvr; pTA131-based deletion vector for HVO_B0235
This study
27
Table 2-3. List of primers used in this study.
Primer Sequence (5'-3')
moaE 447 UP FW CGTGACCGCATCTTAAGGGT
moaE 369 DN RV GTACTGCTTGAGGCGGTCTT
moaE 229 INT FW TACGACTACGCGCTCCTCTC
moaE 750 INT RV AATCGGCACCTCGTCTTTCA
dmsA pKO UP CTAGTGGATCCGACGGCGAGTGTATCG
dmsA pKO DN CGATCAAGCTTGTCACCGAAGATGCGGG
dmsA INT FW GCCACCGAACCGGCGTCGAG
dmsA INT RV AATAAGCTTGTCACCTCCCGCCCCGG
dmsA UP INV TCCGTCGTCACTCATCGAAC
dmsA DN INV GGGAGGTGACTGACCATGAC
HVO_0671 pKO UP CGAATAAGCTTGACGAGATTTACGAGCC
HVO_0671 pKO DN CTAGTGGATCCGTCGAAGTCGGCTATTC
HVO_1471 pKO UP CGATCAAGCTTCGAGAAGTGGTCCTTGA
HVO_1471 pKO DN CTAGTGGATCCGAAGGCGACCTGCAC
HVO_B0367 pKO UP CTGTTAAGCTTGGTGAACTGTCGGCGCTTTTC
HVO_B0367 pKO DN CAACAGGATCCACGACGGTACAGGGCGTGAAG
HVO_B0164 pKO UP CGAATAAGCTTGACTACCACGACGAATC
HVO_B0164 pKO DN CTAGTGGATCCCTCGCGCTTGTAGATG
HVO_0935 pKO UP CGAATAAGCTTGAAGAGGATGAGCAGGA
HVO_0935 pKO DN CTACAGGATCCACGTCCCACTCGGATA
HVO_1908 pKO UP CGATCAAGCTTAGAAGACGCGTCGATAC
HVO_1908 pKO DN CTAGAGGATCCATAGTGTCGGTGCAGG
HVO_B0235 pKO UP GACATGAATTCCCCAGAGGGCACCGATAGAG
HVO_B0235 pKO DN GGTAAGGATCCAGTTGCCAGAGAATAGACACGG
HVO_0671 UP INV AGAAGTGATAGCGTTTCGAGCG
HVO_0671 DN INV TTCCAGTGGGTCGATGTGGTC
HVO_1471 UP INV CCGACGCACCTATCACTAACGA
HVO_1471 DN INV AGCTGAGCTGAGTTTTCGGTCC
HVO_B0367 UP INV TGCATGCGTACTCACTACACCA
HVO_B0367 DN INV ATGACGCGAGAGAGACAGAACC
28
Table 2-3. Continued
Primer Sequence (5'-3')
HVO_B0164 UP INV AAAAGACGGTCGTTAGTGCCAG
HVO_B0164 DN INV GGTGATGCCGAATGAGCACC
HVO_0935 UP INV CGCTGACACACCACACACATAG
HVO_0935 DN INV GATGACTGACTCGTCGGCGTC
HVO_1908 UP INV ATATACCCCGTGCCGTTCGTG
HVO_1908 DN INV GTGAGCGACTGATGGCGTTC
HVO_B0235 UP INV TTTGTCAGAACAGGTGCCGCTC
HVO_B0235 DN INV ACGTTTCGCTGTACTCCTCTCC
29
CHAPTER 4 RESULTS AND DISCUSSION
Deletion of moaE
In order to disrupt molybdopterin synthesis in the methionine auxotroph msrA
msrB mutant (XF130), the moaE gene was deleted from XF130. See methods for
details. Initial pop-out colonies were screened by PCR using primers flanking moaE,
and colonies indicating loss of moaE were streaked for further isolation. This strain
(ΔmetE1/2 ΔmsrA ΔmsrB ΔmoaE) was putatively named ZA106. Further isolated
colonies of ZA106 were then screened using one primer pair flanking moaE and one
pair specific for moaE itself (Figure 3-1). When screened using flanking primers, the
PCR products generated from the deletion mutants were roughly 800 bp less than the
wild-type controls, suggesting that moaE was no longer present. Some minor PCR
products were observed when isolates were screened with moaE specific primers,
however these were expected to be nonspecific products based on size and
comparison to wild type. An isolate (ZA106) that did not display evidence for carrying
the moaE gene based on this PCR analysis (lane 1) was chosen for subsequent strain
preservation and growth assays.
ZA106 Fails to Utilize Methionine Sulfoxide
The isolate ZA106 (ΔmetE1/2 ΔmsrA ΔmsrB ΔmoaE) was assayed for growth
under conditions where MetSO was the only source of methionine present. Growth of
ZA106 was not supported when cultured in GMM supplemented with 0.1 mM MetSO
(Figure 3-2). We note that 0.1 mM Met supplementation was only sufficient to restore
growth of the mutant strains to about half that of the wild type strain H26. This finding
indicated that a higher level of Met supplementation is required under these growth
30
conditions for Met auxotrophs to grow similar to wild type and provided room for
improvement in design of future assays. Still, ZA106 grew similar to the other mutant
strains under Met supplementation, suggested that deletion of moaE did not result in
any significant general impact of growth. Rather, the deletion of moaE was responsible
for loss of residual Msr activity in the cell that previously allowed for growth on MetSO in
strain XF130. Complementation of moaE via plasmid will be completed to confirm this
finding.
Deletion of dmsA
The finding that disruption of molybdopterin synthesis in a methionine auxotroph
with all known Msrs deleted abolished growth on MetSO established clear support that
any major residual Msr activity yet to be identified in H. volcanii was likely molybdopterin
dependent. We next set out to identify what specific molybdoenzyme(s) might be
responsible for this residual Msr activity. Given that moaE deletion in H. volcanii was
previously observed to abolish DMSO reductase activity33, and that DMSO reductases
are known to be capable of reducing MetSO, we chose to delete the gene encoding for
the active site subunit of DMSO reductase—dmsA. Plasmid pJAM114 was introduced in
XF130 (ΔmetE1/2 ΔmsrA ΔmsrB) for deletion of dmsA from the chromosome, as
described in the methods. Candidate deletion mutants were identified by PCR screening
and the strain was putatively named ZA109 (ΔmetE1/2 ΔmsrA ΔmsrB ΔdmsA).
Putative strains were subjected to further isolation and verification of gene loss
by PCR screening. When primers flanking dmsA were used for screening, a PCR
product in the mutants was observed that migrated approximately 1500 bp less than the
product observed in the wild-type control (Figure 3-3). This difference was expected to
be about 2500 bp given the size of dmsA (2532 bp). However, the smaller observed
31
difference can be explained by the migration of the wild-type product some 1000 bp
lower than expected, likely due to incomplete product formation. The mutant product
appeared to be the same size as that of the deletion plasmid itself (lane 14), the fidelity
of which had been verified by DNA sequencing. Furthermore, screening with primers
specific for dmsA over a range of temperatures also suggested that dmsA was no
longer present in ZA109.
ZA109 Retains Ability to Grow on Methionine Sulfoxide
Following generation of strain ZA109, growth was assayed in a similar fashion as
the previous experiment. While ZA106 (ΔmetE1/2 ΔmsrA ΔmsrB ΔmoaE) again failed to
grow on MetSO, ZA109 (ΔmetE1/2 ΔmsrA ΔmsrB ΔdmsA) grew similarly to the wild
type under MetSO supplementation (Figure 3-4). This result suggested that DMSO
reductase was not responsible—or at least not alone—for the residual Msr activity in H.
volcanii. Note that after about 24 h incubation, the OD600 readings for mutant strains
growing without Met appeared to increase slightly but steadily. This increase can also
be observed for strain ZA106 grown on MetSO supplementation. However, this modest
increase can likely be attributed to failure to replenish the water in the outside wells of
the plate, which we have observed results in evaporation of sample cultures and can
impact OD readings. Separate assays conducted in test tubes confirmed that ZA109
was still able to utilize MetSO, whereas ZA106 could not (data not shown).
Identification and Deletion of Additional Candidates
We next sought to test a complete set of candidate ORFs for residual Msr
activity. Seven genes encoding for predicted MPT oxidoreductases were identified and
a summary of their information is shown in Figure 3-5. All seven of these genes were
32
deleted from XF130 (ΔmetE1/2 ΔmsrA ΔmsrB) using the previously described method
and screened for confirmation. Pop-out colonies were screened by the same 500 bp
flanking primers used in construction of the pre-deletion plasmid, and colonies yielding a
PCR product of roughly 1000 bp in the absence of wild-type PCR product were selected
for growth assay as indicated in Figure 3-6. Further isolation and screening of these
isolates remains to be completed. The strains were tentatively named ZA110-116.
Plate Assay Using Candidate Gene Deletion Strains
The isolates of strains ZA110-116 obtained above, in addition to ZA106
(ΔmetE1/2 ΔmsrA ΔmsrB ΔmoaE), were inoculated on Hv-Min plates supplemented
with either 0.25 mM Met or MetSO. After 42 h incubation, all strains were observed to
grow when supplemented with 0.25 mM Met (Figure 3-7). However, on the plate
supplemented with MetSO, minimal to no growth was observed for all strains.
Therefore, incubation of the MetSO supplementation plate was extended to 9 days. By
this point in time, growth was visible for all of the tested strains, but still below the level
observed with Met supplementation. This finding may be explained by the level of
MetSO supplementation not being adequate to achieve the same rate of growth
observed under Met supplementation, or another factor resulting from conducting the
assay on agar plates as opposed to liquid.
More importantly, ZA106 was observed to grow on MetSO supplementation,
which contrasts with the results of both growth assays in liquid media. This level of
growth did at least appear to be the lowest of all strains examined. Interestingly, ZA111
(ΔmetE1/2 ΔmsrA ΔmsrB ΔHVO_1471) appeared to grow at a level similar to ZA106
under MetSO supplementation. This finding suggested that HVO_1471 might be
responsible for at least some of the residual Msr activity present in H. volcanii. Thus, the
33
finding that ZA106 grew on the MetSO supplementation plate presents a double-edged
sword. On one side, the growth of ZA106 is concerning given that growth assays in
liquid indicate the strain is unable to utilize MetSO. On the other, the similar level of
growth observed for ZA111 suggests that HVO_1471 may contain Msr activity.
Use of the plate assay, while intended to provide a quick initial screen of the
candidate Msr deletion strains, did introduce some complications. However,
troubleshooting is to be expected when developing new assays. It may be that ZA106
can utilize MetSO when cultured in a less aerobic environment such as on agar plates.
Alternatively, MetSO may be reduced by other strains on the plate and eventually
diffuse back into the medium, permitting growth of ZA106. Streaking the strains on
separate plate seems to be a good place to start. In any case, further isolation of the
candidate deletion strains and a more thorough PCR screening should be completed.
Troubleshooting of the plate assay should be investigated while also preparing for the
liquid growth assay performed previously. No conclusion should be jumped to
concerning this initial screening of the candidate deletion strains.
34
Figure 4-1. PCR screening confirming deletion of moaE in strain ZA106. Diagram at top
depicts moaE and associated PCR products, to scale. Primers “moaE 447 UP FW” and “moaE 369 DN RV” were used to generate product A (pA), with an expected size in wild-type strains of 1603 bp. Primers “moaE 229 INT FW” and “moaE 750 INT RV” were used to generate product B (pB), with an expected size in wild-type strains of 507 bp. A) Screening of ZA106 isolates vs. wild-type control by primers “moaE 447 UP FW” and “moaE 369 DN RV.” Lane M, marker; 1-12, ZA106 isolates (lysate); 13, wild-type lysate; 14, wild-type genomic DNA extract. B) Screening of ZA106 isolates vs. wild-type control by primers “moaE 229 INT FW” and “moaE 750 INT RV.” Lane M, marker; 1-12, ZA106 isolates (lysate); 13, wild-type lysate; 14, wild-type genomic DNA extract.
35
Figure 4-1. Continued
36
A)
Figure 4-2. Growth assay comparing strains H26, XF127, XF130, and ZA106. Strains
were grown in glycerol minimal media (GMM) supplemented with A) No Met, B) 0.1 mM Met, and C) 0.1 mM MetSO.
37
B)
Figure 4-2. Continued
38
C)
Figure 4-2. Continued
39
Figure 4-3. PCR screening confirming deletion of dmsA in strain ZA109. Diagram at top
depicts dmsA and associated PCR products, to scale. Primers “dmsA pKO UP/DN” were used to generate product A (pA), with an expected size in wild-type strains of 3773 bp. Primers “dmsA INT FW/RV” were used to generate product B (pB), with an expected size in wild-type strains of 2379 bp. A) Screening of ZA109 isolates by primers “dmsA pKO UP/DN.” Lane M, marker; 1-12, ZA109 isolates (lysate); 13, wild-type lysate; 14, dmsA deletion plasmid pJAM3501. Saturated pixels were colored red by image capture software. B) Gradient PCR screening of ZA109 isolates by primers “dmsA INT FW/RV.” Lane M, marker; 1-4, isolate of ZA109; 5-8, additional isolate of ZA109, 9-12, wild-type lysate. Annealing temperatures for each set of samples were 64, 60, 50, and 46 °C, from left to right.
40
Figure 4-3. Continued
41
A)
Figure 4-4. Growth assay comparing strains H26, XF127, XF130, ZA106, and ZA109.
Strains were grown in glycerol minimal media (GMM) supplemented with A) No Met, B) 0.25 mM Met, and C) 0.25 mM MetSO.
42
B)
Figure 4-4. Continued
43
C)
Figure 4-4. Continued
Figure 4-5. Summary of additional candidate gene information and their respective
proteins. Acquired from UniProt database.
44
Figure 4-6. PCR screening of candidate gene deletion isolates for strains ZA110-116.
Primers were the same used in construction of pre-deletion plasmids (pKO UP/DN). Top diagram shows an example gene of 1000 bp, its pKO primer locations, and the respective PCR product (pX) generated. Red boxes denote separate reaction sets for each strain, labels A-G referring to ZA110-116, consecutively. The right most lane in each reaction set is a control using the parent strain DNA as template, yielding a PCR product approximately 1000 bp plus the gene size. Potential deletion isolates display an absence of this PCR product, with a strong product instead at roughly 1000 bp. Candidate deletion isolates marked by red asterisks were selected for growth assay and further isolation.
45
Figure 4-7. Plate assay for growth of additional candidate mutant strains on MetSO.
The plate on the left is supplemented with 0.25 mM Met and was incubated for 42 hours; on right, 0.25 mM MetSO supplementation with 9 days incubation. Wedges A-G are inoculated with the additional candidate mutant strains, wedge Z is inoculated with ZA106.
46
CHAPTER 5 CONCLUSIONS
Observations preceding this study indicated that an H. volcanii methionine
auxotroph with both predicted Msrs (MsrA/B) deleted was still capable of growth on
MetSO as the sole source of methionine. This discovery came as a surprise given that
no additional Msrs are predicted in H. volcanii, nor do they appear to be present in other
Halobacteria genomes. Since many oxidoreductases are molybdopterin dependent, we
deleted a gene thought to be a critical component of the molybdopterin synthesis
machinery in H. volcanii. Successful deletion of moaE revealed that the residual Msr
activity of H. volcanii was indeed molybdopterin dependent, as the strain was no longer
able to utilize MetSO for growth. This finding provided a clear step forward in identifying
the enzyme(s) responsible for residual Msr activity in H. volcanii.
Further investigation found that DMSO reductase is likely not the source of
residual Msr activity. Seven additional candidate genes were deleted from H. volcanii,
but initial experiments have not been conclusive as to whether one or more of these
candidates are culprits. Continuation of the growth assay initiated in this study may
reveal that one of these candidate genes is responsible. Further isolation of strains
ZA110-116 and more thorough confirmation of their deletion should be completed
regardless before these genes are ruled out. Alternatively, other methods may need to
be utilized to identify this unknown source of Msr activity.
Preparation of the individual diastereoisomers Met-(R)-SO and Met-(S)-SO could
provide more information regarding the stereospecificity of the residual Msr activity.
Given that prior work did not observe Msr activity in an H. volcanii ∆msrA ∆msrB mutant
using the peptide mimic dabsyl-MetSO35, it seems that any major residual activity would
47
likely be specific for free MetSO. Should targeted approaches fail, broader genetic or
biochemical approaches such as transposon mutagenesis or cell lysate fractionation
could be utilized to identify the unknown source of Msr activity in H. volcanii. Such an
effort would be helpful in furthering our understanding of Msrs and their peculiar
distribution in archaea.
48
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52
BIOGRAPHICAL SKETCH
Zachary Adams obtained his Bachelor of Science in 2016 from the University of
Florida. During his senior year, he worked in the laboratory of Dr. Julie Maupin-Furlow in
the Microbiology and Cell Science Department. He chose to continue his studies there,
and later obtained his Master of Science degree in the Fall of 2018.