Siderophore Production by Heterotrophic Bacterial Isolates
from the Costa Rica Upwelling Dome
By
Whitney B. Krey
B.S., Texas A&M University, 2005
Submitted in partial fulfillment of the requirements of the degree of
Master of Science
at the
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
and the
WOODS HOLE OCEANOGRAPHIC INSTITUTION
June 2008
02008 Whitney B. Krey
All rights reserved.
The author hereby grants to MIT and WHOI permission to reproduce paper and
electronic copies of this thesis in whole or in part and to distribute them publicly.
MCHIvs
Signature of Author
_
Joint Program in Oce
Massachusetts Institute of Te
_
~phy/Applied OceF) Science and Engineering
y and Woods Hole Oceanographic Institution
June 2008
Certified by
--
Thesis Supervisor and C air, Joint C
Massachusetts Institute of echnolo
Mak Saito
Thesis Supervisor
Ed DeLong
mittee for Biological Oceanography
oods Hole Oceanographic Institution
Abstract:
An increased understanding of heterotrophic bacterial strategies for acquiring nutrients
and trace elements is critical for elucidating their impact on biogeochemical cycling in
the ocean. It is estimated that iron is a limiting nutrient for phytoplankton growth in over
30% of the open ocean, but still little is known about bacterial strategies for iron
acquisition. Siderophore (Fe ligand) production by bacteria may play a major role in
influencing the bioavailability of iron in the ocean. Despite the importance of
siderophores in the environment, only limited information from a select group of bacteria
is available. On a cruise through the Costa Rica Dome (CRD) upwelling region in July
2005, a library of 867 isolates from five depth profiles inside and outside of the dome
was obtained and screened for siderophore production using the Chrome Azurol-S (CAS)
assay. Phylogenetic affiliation of 134 isolates was determined by sequencing the 16s
rDNA gene, and determined that gamma proteobacteria such as Alteromonas,
Pseudoalteromonas,Halomonas,and Marinobacterdominated the collection, while
alpha-proteobacteria such as Roseobacter were also represented. The isolates obtained
from stations in the CRD showed greater siderophore-producing capabilities between
55m and 100m while strains isolated from outside the CRD had shallower peak (-8-35m)
production. Functional group determination showed that hydroxamate production
dominated from 50-150m, while hydroxamate and catechol production is roughly equal
in shallower waters. By characterizing the siderophores produced by these isolates and
determining the genetic make-up of the population, these findings further our
understanding of how heterotrophic microbes affect biogeochemical processes and the
competitive nature of nutrient acquisition.
History and Background:
Microbes and their role in the environment
Microbes are ubiquitous and significant regulators of many of earth's fundamental
processes. Bacteria are found nearly everywhere on Earth, growing in soil, acidic hot
springs, radioactive waste, seawater, and deep in the Earth's crust (Fredrickson et al.
2004). Forty million bacterial cells can be found in a gram of soil, and more than a
million bacterial cells can be found in a milliliter of fresh water. Approximately 5x 1030
bacteria can be found on Earth (Whitman et al. 1998). In addition to their notorious role
of causing disease, bacteria are vital in the recycling of nutrients. Despite their
importance, most of these bacteria have not been characterized, and less than half of the
phyla of bacteria have species that can be cultured in the laboratory (Whitman et al.
1998).
Advances in genomics capabilities are providing a more refined view of the capabilities
and complexity of the previously underestimated oceanic microbial community,
particularly with respect to diversity and functionality. The challenge is integrating these
discoveries at the systems level to elucidate specific microbial functional roles and
overall system resilience. Understanding the role of microbes in structuring healthy and
stressed marine ecosystems will provide the mechanistic basis for extrapolative models.
Genome sequencing has confronted scientists with the reality that many functional
capabilities and molecular mechanisms remain unknown. Evidence suggests that
important discoveries remain. The relatively recent discovery of proteorhodopsin genes
in bacteria is an example. This light-driven pigment converts light energy into an
electrical gradient across the cell membrane, and is thought to be an additional, but not
primary, source of energy (Beja et al. 2001). Previously thought to only exist in Archaea,
proteorhodopsin has now been found in divergent marine bacterial taxa in diverse
environments. Another example of an important metabolic discovery is that of the
oceanwide distribution of anoxygenic phototrophic bacteria (Madigan 2003). Their
contribution to oceanic energy and carbon budgets is much larger than previously
thought. Other biogeochemical budgets, such as nitrogen, must now be reconsidered due
to fresh evidence for nitrogen fixation in common oceanic systems. It is possible that
these, as well as undiscovered metabolic pathways, contribute to oceanic biogeochemical
budgets in ways that change our current estimates.
Biological importance of iron
These notable discoveries highlight our current knowledge limitations, as well as the
importance of a thorough understanding of microbial metabolism and environmental
interactions. A key component needed to link the contribution of bacteria to oceanic
nutrient cycles and higher trophic levels is knowledge of the uptake strategies for
nutrients that limit growth (Martinez et al. 2000). One such nutrient is iron. Along with
other necessary nutrients such as carbon, nitrogen, and phosphorous, microbes need to
take up iron. On a broad scale, iron's importance is illustrated by its involvement with
regulating coastal and open ocean microbial diversity, controlling primary productivity
levels in large areas of the open ocean, and controlling phytoplankton populations in
upwelling regions (Ryther and Kramer 1961, Kirchman et al. 2000, Barber and Ryther
1969). On the cellular level, iron is involved in important biological processes such as
photosynthesis, nitrogen fixation, methanogenesis, H2 production, the trichloroacetic acid
(TCA) cycle, oxygen transport, gene regulation, DNA biosynthesis/repair, and
detoxification of free radicals (Andrews et al 2003, Crichton and Ward 1992, 1998).
Oceanic algae and bacteria have a high rate of division and turnover relative to land
plants, creating a high turnover of nutrients. Biologically available iron is almost
completely depleted in surface waters due to this high turnover and cellular quota.
Surface iron concentrations are approximately one millionth that of the concentration
inside of the microbial cells (Morel and Price 2003). While most nutrients are effectively
recycled and regenerated in their respective part of the water column, depletion due to
export is a problem; abiotic scavenging and sinking particles are constantly depleting the
iron pool. Because of these export factors, water column iron concentrations increase
slightly with depth due to remineralization by bacteria below the photic zone.
Iron has two stable oxidation states (Fe (II) and Fe (III)) that can be manipulated to
catalyze a wide number of biochemical reactions. For these biochemical reactions, iron is
used by incorporation into proteins, or in complexes forming part of iron-sulfur clusters
or heme groups. Incorporating iron into proteins allows for control of the local redox
potential (ranging from -300 to +700mV), geometry and spin state of the iron atoms, such
that they can fulfill their necessary biological function (Andrews 2003). While Fe (III) is
the most widely available form of iron in the ocean, Fe (II) is ultimately the form that
biology utilizes. While certain transporters and iron reductases can draw Fe (II) into the
cell, known siderophores only bind Fe (III). Internal mechanisms then reduce the Fe (III)
to Fe (II), allowing it to be used within the cell.
The versatility of iron makes it biologically useful, but there are downsides to using this
element. Although iron is the 4t h most abundant element in earth's crust, it is widely
unavailable to biological systems in the ocean due to its insolubility in aerobic
environments at neutral pH, as well as biological factors such as competitive uptake
(Sunda and Huntsman 1995). In all, it is estimated that the growth of 30% of the primary
production in the open ocean is limited by iron (Martin et al. 1994, Coale et al. 1996,
Moore et al. 2004). Iron in surface seawater is present in extremely low concentrations
[typically from 20 pM to 1 nM], which limits primary production by phytoplankton in
regions characterized by high concentrations of nitrate and other nutrients but lower than
expected concentrations of chlorophyll (HNLC, high nitrate low chlorophyll) (Gordon et
al 1982, Landing and Bruland 1987, Martin at al. 1989, 1991, Bruland et al. 1994).
HNLC regions include the subarctic Pacific, eastern equatorial Pacific, the Southern
Ocean, and upwelling regions (de Baar et al. 1990, Martin and Fitzwater 1990, Coale et
al. 1996). Heterotrophic bacteria are also limited by low iron levels in HNLC regions,
though the complexity of their iron requirements and uptake strategies are poorly
understood. In regions where phytoplankton are clearly iron limited, conflicting reports
exist on iron limitation in heterotrophic bacteria, further highlighting our limited
understanding of the complex cycling of this essential nutrient (Kirchman et al. 2000).
It is estimated that >99% of the dissolved iron in seawater is complexed by organic
ligands in the North Sea, Western Mediterranean, North Pacific, Northwest Atlantic, and
equatorial Pacific (Gledhill and van den Berg 1994, van den Berg 1995, Rue and Bruland
1995, Wu and Luther 1995, Rue and Bruland 19971). The structural composition of this
organic ligand pool is largely unknown. One major stumbling block in the
characterization of the natural organic ligand pool is the large-scale sampling that is
required. A milligram or more of ligand may be required for characterization, limiting the
resolution of the samples that can be analyzed (Macrellis et al. 2001). While no
siderophore structures have been characterized from the field, lab and field experiments
show that siderophores are a component of this pool (Haygood et al. 1993, Rue and
Bruland 1995, Lewis et al. 1995, Wilhelm et al. 1998, Hudson 1998, Hutchins et al.
1999b). Organic ligands isolated in the California upwelling region have been shown to
possess iron-binding functional groups typical of siderophores (Macrellis et al. 2001).
Most other thorough siderophore characterization has taken place on marine bacterial
isolates in the laboratory, allowing for intense structural and binding constant
determination. While it has been shown that many bacteria are capable of producing
organic iron ligands, and that such ligands exist in the field, the link between the two is
still largely a mystery. We need to know more about the source, characteristics and fate
of these ubiquitous Fe-binding compounds.
Iron acquisition strategies
Heterotrophic bacteria constitute up to half of the total particulate organic carbon in
ocean waters, and in some regions, such as the subarctic Pacific, heterotrophic bacteria
can even contain higher cellular concentrations of iron than phytoplankton. Heterotrophic
bacteria thus compete successfully for iron against phytoplankton and cyanobacteria and
play a substantial role in the biogeochemical cycling of iron in the ocean. However, little
is known about the molecular mechanisms used by marine bacteria to sequester iron.
Heterotrophic bacteria have evolved Fe (III) transport systems that enable them to grow
in environments containing extraordinarily low concentrations of iron. When iron is
scarce, many of these organisms excrete low-molecular mass (300-1000 Da) Fe-binding
chelators, called siderophores, that bind free Fe (III) (Neilands and Nakamura 2001).
Siderophore production has also been shown in cyanobacteria (Armstrong and van
Baalen 1979, Wilhelm and Trick 1994), algae, fungi and plants (Neilands 1984, Suguira
and Nomoto 1984). Some marine heterotrophs produce and/or excrete siderophores
during iron stress and acquire siderophore-bound iron to fulfill their iron requirements
(Trick and Wilhelm 1995).
Many types of siderophores have been isolated from marine bacterial strains.
Hydroxamate and catecholate type siderophores seem to be the most prevalent in the
bacterial strains studied, but other functional groups such as alpha-hydroxy acids,
carboxylic acids, and 2-hydroxyphenyl oxazoline are also found (Hider 1984). However,
many of the structures have not undergone full characterization (Lewis et al. 1995). A
handful of complete siderophore structures have been determined, but these were isolated
from a very small portion of the ocean's diverse bacterial community - and only
represent structures produced under laboratory conditions. This highlights the need to
better link bacterial siderophore production with behavior in the field.
The connection between iron transport and siderophores in marine microbes is less
understood in comparison to the terrestrial model organisms. Terrestrial siderophores
such as enterobactin and desferrioxamine B are produced by E. coli and Streptomyces
(O'Brien and Gibson 1970, Pollack and Neilands 1970, Bickel et al. 1960). The precise
molecular mechanisms of Fe-siderophore transport are now being elucidated in E. coli
and other laboratory strains (Ferguson et al. 1998; Buchanan et al. 1999; Ratledge and
Dover 2000). In these model organisms, the Fe-siderophore complex attaches to its
cognate receptor on the outer membrane of the cell and is subsequently internalized
(Braun and Killman 1999; Ratledge and Dover 2000). Some bacterial receptors recognize
more than one siderophore, not necessarily of the same structural type (e.g., FhuE of E.
coli) (van der Helm 1998). Many bacteria express receptors for siderophores released by
other species or for Fe sources contained in their hosts (Braun et al. 1998; Ratledge and
Dover 2000). Such uptake systems provide a high degree of selectivity and regulation and
enable microbes to scavenge and solubilize Fe from otherwise inaccessible sources.
Much less is known about iron transport by aquatic bacteria, particularly among the
heterotrophic marine species. Short-term uptake experiments show that bacteria take up
most of the dissolved iron (Tortell et al. 1996; Maldonado and Price 1999) and contain
relatively large amounts of Fe in their biomass compared to other living organisms
(Tortell et al. 1996). The reasons for the success of bacteria in acquiring iron under these
conditions are not well understood, but they may have to do with their ability to produce
siderophores and to transport ferric siderophore complexes. The ability of heterotrophic
bacteria to produce siderophores has been well established in the laboratory (Barbeau et
al. 2003, Granger and Price 1999, Martinez et al. 2001, 2003), and the presence of
siderophore-like iron ligands has been established in seawater. A more thorough
understanding of the molecular mechanics behind siderophore production will hopefully
lead to the ability to actively detect their production in the field, and thus help elucidate
the impact heterotrophic bacteria have on the iron cycle in the ocean.
Study site: The Costa Rica upwelling dome
The Costa Rica Dome (CRD) is an upwelling region located in the eastern tropical
Pacific Ocean close to 8"N, 90'W. It is a seasonal phenomenon, shifting shape, size, and
location as wind stress changes. The thermocline of this area reaches to approximately 10
m, creating a dome-like shape and thus giving the feature its name. The Costa Rica
Coastal Current, Eastern Equatorial Current, and North Equatorial Current all converge in
this area to create a cyclonic gyre. Upwelling velocities as high as 10-4 cm/sec have been
calculated (Hofmann et al. 1981). Upwelling water appears to originate from between 75
and 200 m. (Broenkow 1965).
Upwelling is a phenomenon that brings cold, nutrient rich water up from depth, typically
causing increased primary productivity. Upwelling regions are generally dominated by
diatom blooms at the autotrophic level. However, the CRD is unique in that it is
dominated by cyanobacteria - specifically Synechococcus. Cell densities of
Synechococcus are higher in the CRD than any other open ocean environment measured,
ranging from 0.5x10 6 to 1.5x10 6 cells/ml (Goericke and Welschmeyer 1993, Campbell et
al. 1994; DuRand et al. 2001, Saito et al. 2005). These numbers are approximately an
order of magnitude larger than typical cyanobacterial concentrations in the open ocean
(Saito et al. 2005). The KN182-50 cruise in July 2005 was dedicated to characterizing
trace metals and cyanobacterial population dynamics in and outside of the CRD.
The heterotrophic bacteria in this distinctive regime are largely uncharacterized. This
thesis work focuses on the phylogeny of heterotrophic bacterial isolates from the CRD
and their siderophore-producing capabilities with the goal of gaining greater insight into
the contribution of heterotrophs to oceanic iron cycling.
Methods:
Cruise Sampling: Samples used in this work were obtained aboard the UNOLS ship R/V
Knorr, July 15-August 2, 2005 on the KN182-50 cruise.
Cruise track
10
.,q
-100
-98
-96
-94
92
-90
-88
-86
-84
.82
-80
Figure 1: KN182-50 cruise track. Stations where culturing took place are labeled with
yellow circles. Station 17 was located outside of the upwelling dome in a predicted high
nutrient, low chlorophyll (HNLC) region.
Culturing: Water samples were plated on 25 mm Petri dishes containing complex agar
media from the five stations indicated on the cruise track (Fig. 1). Samples were taken at
five depths, ranging from 8-200 m following the water column features. Samples were
diluted 10- 1, 10-2, and 10-3 with sterile autoclaved seawater in addition to the undiluted
sample. 20 gl of each dilution was plated in triplicate onto the complex agar media and
Chrome Azurol-S (CAS) agar media. Cultures were incubated at room temperature.
Batch mediapreparation: 1.0 g yeast extract (Difco), 1.0 g tryptone (Difco), 24.7 g
NaCl, 0.7 g KC1, 6.3 g MgSO 4 -7H2 0, 4.6 g MgC12-6H 20, 1.2 g CaCl 2 -2H20, 0.2 g
NaHCO 3 , 15 g agar (Difco) per liter (Wagner-Dobler et. al 2003). Chemicals and agar
were added to Milli-Q water and then heat sterilized. CAS reagents were 100 ml diH 20,
60.5 mg chrome azurol S, 1 ml Ironll-KCl (10x) solution (10X solution is 10 mM
FeC13-6H 20 and 100 mM KC1), and 72.9 mg hexadecyltrimethylammonium bromide in
1L sterile seawater (Schwyn and Neilands 1987). Samples were incubated at room
temperature for three days before storage at 4°C. Culture plates were shipped back to the
laboratory immediately after the cruise.
Six representatives of each colony morphology from cultured stations at each depth and
dilution were picked and transferred to 150 jil of liquid media in sterile 96-well plates
(above recipe excluding agar and CAS reagents). 960 colonies were picked, of which 867
were ultimately isolated and survived. After three days of growth, cultures were streaked
to isolation three times in sterile petri dishes containing the solid version of the same
media. Isolated colonies were then picked and transferred to the liquid version of their
initial isolation media (CAS was excluded from the recipe at this stage). Cultures were
maintained in 96-well plate format, 150 tl media/well. Every 1.5 weeks 3 ptl of each
culture was transferred to fresh media using aseptic techniques. After initial chemical
assays, the cultures were frozen in duplicate at -800 C with 5% DMSO for preservation.
DNA Extractions:The cultures were diluted 1:10 culture:sterile water in fresh, sterile 96well plates (pre-DMSO preservation). The samples were run in an Eppendorf
Mastercycler thermocycler for 10 mins at 95 0 C. The samples were then centrifuged for 5
min at 3000 x g to pellet cellular debris. Plates were sealed and stored in a -20 0 C freezer.
PCR: DNA was amplified using 8f and 1492r bacterial 16S rDNA primers: 8f: 5'AGAGTTTGATCCTGGCTCAG-3', and 1492r: 5'-GGTTACCTTGTTACGACTT-3'.
5 ptl of DNA template was used in a total reaction volume of 27.5 p1. Samples were
amplified in 96-well format. The amplification program was 95 0 C for 3 min, followed by
25 cycles of: 95 0 C 1 min, 60 0 C 1 min, 720 C 1 min, and final extension of 72 0 C for 10
min. 1%agarose gel electrophoresis was used to confirm presence and quality of
extracted DNA.
Restrictionfragment length polymorphism (RFLP)binning: In an effort to screen the
867 culture isolates obtained on the cruise, RFLP was performed as a preliminary
diversity screen. Each PCR product was digested in a mixture containing 0.5 pl of 20,000
U/ml TaqI restriction endonuclease (New England Biolabs, Beverly, MA), 1 [l of 10X
Taql buffer, 0.1 gl of 100ug/ml bovine serum albumin, 3.4 gl of ddH20, and 5 Rl of PCR
product. The digests were incubated in an Eppendorf Mastercycler at 65"C for 1 hr. RFLP
digest patterns were visualized with 2% agarose gel electrophoresis (Ehrenreich et al.
2005). A minimum of three representatives from each RFLP pattern obtained were
chosen for further phylogenetic analysis.
PCR purification: Samples chosen for sequencing were then purified using the Quiagen
PCR Purification kit according to the manufacturer's protocol.
Sequencing: Sequencing was carried out by Northwoods DNA, Inc. (Solway, MN). The
16S rDNA PCR product obtained were used for sequencing. Sequencing primers used
were: 8f: 5'-AGAGTTTGATCCTGGCTCAG-3', 926r: 5'-ACCGCTTGTGCGGGCCC3', 519r: 5'-GWATTACCGCGGCKGCTG-3', 704r: 5'-GTAGCGGTGAAATGCGTAG
A-3', and 1492r: 5'-GGTTACCTTGTTACGACTT-3'. High quality DNA sequences
were obtained.
PhylogeneticAnalysis: Sequence assimilation was conducted in Sequencher. Sequences
were overlapped and only clean double-stranded sequences in excess of 1400 bp were
used in the alignment. Alignments and tree-building were completed in ARB using the
GreenGenes database. The most up-to-date version of the GreenGenes database was used
in tree building. All reference sequence alignments were double-checked for high quality
and accurate alignments. X-Fig and PAUP were used for further tree modifications
including creation of neighbor-joining trees and bootstrapping. Parsimony and distance
bootstrapping/distance calculations all carried out at 1000x runs. Depth and siderophore
data were added to trees in Adobe Illustrator CS.
Chrome Azurol-S (CAS) assay: All isolates were inoculated in triplicate in complex
media with Dipyridyl added (200 iM final concentration) and iron-replete media.
Original isolation media was used. Uninoculated dilution series of media with Desferral
and Dipyridyl were used as positive and negative controls, respectively. All cultures were
allowed to grow for 24 h at room temperature. At timepoint, samples were spun down for
5 minutes at 3000 x g to pellet cells, and the supernatant was transferred to sterile 96-well
plates. CAS reagent was added (15 gl per 150 gl media). Samples were allowed to react
for 2 h before photographs of plates were taken and subsequent analysis of the assay
began.
After binning via RFLP, the samples chosen for sequencing were analyzed again for CAS
reactivity and siderophore structure type (using Arnow and Czacky assays below).
Samples were CAS tested using the same procedure as above.
Siderophore category determination:
The cultures that were chosen for sequencing were then grown up in triplicate 5ml
volumes of iron replete and dipyridyl-spiked media using 50 ýtl initial culture. Triplicate
uninnoculated aliquots of iron replete and dipyridyl-spiked media were also incubated.
After 24 h of growth at room temperature, samples were spun down and 1 ml supernatant
was transferred to a clean test tube, respectively, for each assay below.
Arnow assay:
This assay is the established and accepted assay use to test for the presence of catecholtype siderophores (among other chemical properties not relevant in this study).
One ml of each sample was placed in individual test-tubes graduate at 5 ml. One ml of
the standard solution (50 mg 3,4-dihydroxyphenylalanine in 500 ml distilled water, add 2
ml 0.1 N hydrochloric acid and bring to one liter volume with distilled water) was also
placed in a graduated test tube. To each test tube the following was added in the given
order, swirling after each addition: 1 ml 0.5 N hydrochloric acid, 1 ml nitrate-molybdate
reagent (10 g sodium nitrate and 10 g sodium molybdate in 100 ml distilled water), 1 ml
1 N sodium hydroxide, and distilled water to bring the volume up to 5 ml (Arnow 1937).
Czsaky assay:
This assay is the established and accepted assay used to test for the presence of free
hydoxamates, or hydroxyl-type siderophores.
To make standards, 1-5 ml of 0.2-0.5 mg hydroxylamine N is placed in a 10 ml calibrated
tube and 1 ml of sulphanilic acid (10 g sulphanilic acid dissolved in 1000 ml of 30%
acetic acid by heating in a water bath) and 0.5 ml iodine solution (1.3 g iodine in 100ml
glacial acetic acid) measured to it. After 3-5 minutes the excess of iodine is destroyed
with I ml of sodium arsenite solution (2 g sodium arsenite in 100 ml distilled water) after
adding 1 ml alpha-naphthylamine solution (3 g alpha-naphthyline dissolved in 1000 ml of
30% acetic acid.) the mixture is made up with distilled water to 10 ml. After 20-30
minutes, the color is developed.
A sample of 1ml of the solution to be tested and 1 ml of 6 N sulfuric acid was placed in a
Pyrex glass test tube with ground-in stopper carrying capillary tubing. The solution was
boiled in a water bath for 6 hours and transferred to a 10ml tube. The excess of sulfuric
acid was buffered with 3ml of 35% sodium acetate, and the free hydroxylamine was
detected using the method above (Czsaky 1948).
Along with the samples, negative controls of uninnoculated iron replete and dipyridylspiked media were tested for the presence of hydroxamates.
Results:
Siderophore Production:
All 867 isolates were tested for siderophore production. 340 out of the 867 (39.2%) tested
positive for siderophore production. 61 of the 340 have sequenced representatives in this
study. Results organized by depth and station can be found below in Figures 2-6. Stations
2, 5, 7, 15 and 17 are represented.
Station 2 Siderophore Producing Isolates by Depth
8m
15 m
35 m
55 m
Ill~llll
I1ll~ll~llll///II
-
i
I
] . I............
l • l...
mH/l
l /il •
100 m
-i
150m
0%
n
ii~l~[rr
20%
40%
60%
80%
100%
Percent Total Isolates
Figure 2: Station 2 siderophore producing isolates by depth. Black shows siderophore
producers, gray shows non-producers. Shallow depth isolates failed to produce
siderophores under experimental conditions. Siderophore producing isolates appeared at
35m and peaked at a depth of 55m.
Station 5 Siderophore Producing Isolates by Depth
8m
inmm~mmrni,
15 m
35 m
-----
55 m
----
100 m
150m
0%
10%
20%
30%
40%
50%
60%
70%
80%
90%
100%
Percent Total Isolates
Figure 3: Station 5 siderophore producing isolates by depth. Black shows siderophore
producers, gray shows non-producers. All depths showed isolate siderophore production
under experimental conditions. Production peaked at a slightly deeper depth, 100m at this
station versus 55m at Station 2.
Station 7 Siderophore Producing Isolates by Depth
-
-
0)%
20%
8m
15 m
35 m
55 m
100 m
150m
-
--
-
iniin
iniin
-
*i
40%
60%
Percent Total Isolates
80%
100%
Figure 4: Station 7 siderophore producing isolates by depth. Black shows siderophore
producers, gray shows non-producers. All depths showed relatively even numbers of
siderophore producing isolates with the exception of a production peak at 100m.
Station 15 Siderophore Producing Isolates by Depth
8m
15 m
35 m
-....-
N..
-
-. ...
55 m
100 m
150m
0)%
10%
20%
30%
40%
50%
60%
Percent Total Isolates
70%
80%
90%
100%
Figure 5: Station 15 siderophore producing isolates by depth. Black shows
siderophore producers, gray shows non-producers. Siderophore producing isolates have a
major peak at the shallower depth of 35m and another minor peak at 150m. All depths
contained siderophore producing isolates.
Station 17 Siderophore Producing Isolates by Depth
8m
15m
E 35m
55m
150m
0%
10%
20%
30%
40%
50%
60%
70%
80%
90%
100%
Percent Total Isolates
Figure 6: Station 17 siderophore producing isolates by depth. Black shows
siderophore producers, gray shows non-producers. This station showed the highest
fraction of isolates having siderophore production capabilities in the shallow 8m and 15m
waters.
Siderophore production capabilities appeared to increase with depth in stations 2-7. This
trend appeared to reverse in stations 15 and 17, with production peaking at shallower
depths. All stations showed siderophore production at all depths sampled with the
exception of station 2, which did not show production capabilities at 8m or 35m under the
experimental conditions. All stations contained at least one depth where a majority of
isolates had siderophore producing capabilities.
Siderophore structural determination:
Hydroxamate vs. catecholate structure was determined for all sequenced isolates. Results
can be found in Figure 7 for overall siderophore type vs. depth, and Figures 15 and 16 for
the same data for stations 2-15 and 17, respectively. Data for isolates from all stations
shows approximately even hydroxamate vs catechol production in shallow waters with
more isolates producing hydroxamates than catechols at 100m and 150m. The
siderophore structure type of three isolates was not determined due to unreactiveness with
both the Czsaky and Arnow assays.
Siderophore type by depth, all stations
8m
35m
O
' Indeterminate
!
50m
Hydroxamate
U Catechol
100m
150m
0
2
4
6
8
10
12
14
# isolates
Figure 7: Siderophore type by depth, all stations.
When station 17 is omitted from the overall analysis, hydroxamate-type siderophores
appear to dominate slightly more at depth (Figure 8). Station 17 has an equal number of
hydroxamate and catechol producers at 8m and 35m, but hydroxamates have a majority at
55m and 100m (Figure 9). Only catechols were detected at 150m.
Siderophore type by depth, Stations 2,5, 7, 15
8m
35m
II·_
I
/1
I
IXNX
e
3
150m
C Indeterminate
N Hydroxamate
I Catechol
II
50m
6
# isolates
Figure 8: Siderophore type by depth, stations 2, 5, 7, 15.
Siderophore type by depth, Station 17
8m
35m
Indeterminate
•IN
(
50m
lfa
U Hydroxamate
Catechol
150m
0
0.5
1
1.5
# isolates
Figure 9: Siderophore type by depth, station 17.
2
2.5
Alteromonas and Pseudoalteromonaswere the most numerous groups within the isolates
chosen for sequencing. Figures 10 and 11 show their siderophore production versus depth
for all stations.
Hydroxamate vs. Catechol production in Alteromonas
8m
35m
50m
N
n Catechol
N Hydroxamate
N
100m
150m
0
1
2
3
4
5
6
# sequenced isolates
Figure 10: Alteromonas siderophore production vs. depth. Catechol production
occurs in shallow isolates only. Hydroxamate production capabilities increase at depth.
Hydroxamate vs. Catechol production in Pseudoalteromonas
8m
35m
MCatechol
100m
150m
0
1
2
3
4
# sequenced isolates
Figure 11: Pseudoalteromonas siderophore production vs. depth. Catechol production
present at all depths. Hydroxamate production is only present at two depths. Catechol
production is dominant overall for this subset of this taxonomic group.
16S rDNA Phylogeny:
The sequenced isolates were predominantly gamma and alpha proteobacteria, with a few
representatives from Flavobacterialesand Actinomycinae. Out of the 134 isolates from
which high quality 16s rDNA sequences were obtained, 44 were Alteromonas, 37 were
Pseudoalteromonas,27 were Halomonas, 8 were Marinobacter,6 were Idiomarina,5
were Cytophaga, 3 were Roseobacter, 2 were Porphyrobacter,one was Alcanivoras, and
one was an Aureo-Microbacterium.Figure 12 depicts the overall relationship of all
sequenced isolates with each other in relation to the major bacterial, archaeal, and
eukaryotic taxonomic groups.
Bacteroiakles
E
Flavobacterlales
Cyanobacterla
la
Archaea
s
0.10
larla
GammaproMobeclarla
Figure 12: Overall relationship of cultured groups to other major taxonomic groups.
Isolates from this study are members of red groups.
RFLP revealed 24 patterns within the original 867 isolates. Of these 24 patterns, 76.5%
of isolates fell within 4 of the patterns (32.1%, 25.9%, 11.1%, and 7.2%, see Figure 13).
All other patterns ranged from 0.2%-5.3%.
Percent of isolates per RFLP pattern, all stations
35
30
25
S20
•
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10
5
0
1
3
5
7
9
11
13
15
17
19
21
23
Pattern number
Figure 13: Percent of isolates per RFLP pattern obtained for all stations.
When broken down by phylogeny after 16S rDNA sequencing, Alteromonas was made
up of 9 patterns. Proportions among these 9 patterns very closely followed those of the
overall group of isolates, with a slight increase in the largest group (42.9% in
Alteromonas versus 32.1% overall, see Figure 14). Pseudoalteromonaswas more
scattered, with 11 patterns represented and no group reaching over 23% of the total. Six
of the 11 patterns ranged from 7.4%-22.2% (See Figure 15). Halomonas was composed
of 11 distinct RFLP patterns with a fairly even distribution between 8 of the 12 (12%-
20% of the total, Figure 16). Five patterns make up Marinobacterwith isolates nearly
evenly distributed between them (Figure 17). Three patterns make up the Idiomarina
isolates but with so few isolates, it is difficult to draw conclusions from that data.
Roseobacterand Cytophaga follow that trend with three and four patterns, respectively.
Percent Alteromonas isolates
per RFLP pattern
Percent Pseudoalteromonas
isolates per RFLP pattern
3u
25
45
40
20
35
o 30
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10
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10
5
5
0
n
Individual pattern
Percent Halomonas isolates
per RFLP pattern
Individual pattern
Percent Marinobacter isolates
per RFLP pattern
25
30
20
25
20
o 15
15
5
5
0
Individual pattern
Individual pattern
Figures 14-17: Percent total isolates for Alteromonas, Pseudoalteromonas,
Halomonas, and Marinobacterdistributed by RFLP pattern.
When the siderophore production types are examined, catechol producers are spread
among 13 patterns while hydroxamate producers are spread among 15 patterns. While
both siderophore types basically follow the general pattern proportions of the overall
isolate library, catechol producers have a slightly smaller representation of the largest
pattern group (14.3% versus 32%) that is spread out amongst the other smaller groups.
Both types of producers overlapped on all but three patterns, which each had exclusively
from the other type of siderophore.
Figures 18-24 illustrate the phylogeny of the isolated sequences with siderophore
production, station number, and depth information included.
Figure 18: Phylogenetic tree of cultured Alteromonas isolates. Siderophore
production, type of siderophore produced, depth of isolation, and station are also
indicated. Siderophore production versus depth and station appeared to be fairly random.
No major patterns within the Alteromonas could be distinguished within this set of
isolates.
Phylogeny and siderophore production of Alteromonas isolates
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production capabilities, type of siderophore produced, depth of isolation, and station are
also indicated. Isolates appear to be extremely closely related. Siderophore production
versus depth and station appeared to be fairly random, with the exception of a small
group of isolates from stations 15 and 17 that formed their own small group. No major
patterns within the Pseudoalteromonascould be distinguished within this set of isolates.
Phylogeny and siderophore production of Pseudoalteromones isolates
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production capabilities, type of siderophore produced, depth of isolation, and station are
also indicated. Isolates appear to fall into two distinct phylogenetic groups, although they
are not grouped according to depth, station, or siderophore production.
Phylogeny and siderophore production inMarinobacter isolates
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Figure 21: Phylogenetic tree of cultured Idiomarina isolates. Siderophore production
capabilities, type of siderophore produced, depth of isolation, and station are also
indicated. Within the highest-quality sequences used in building this tree, only stations 2
and 5 had Idiomarinaisolates.
Phylogeny and siderophore production in Idlomarina Isolates
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capabilities, type of siderophore produced, depth of isolation, and station are also
indicated. Siderophore production and type do not coincide with phylogenetic affiliations.
Phylogeny and siderophore production in Halomones isolates
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Figure 23: Phylogenetic tree of cultured Cytophaga isolates. Siderophore production
capabilities, type of siderophore produced, depth of isolation, and station are also
indicated. Three distinctive groups occur phylogenetically, although siderophore
production and type of siderophore do not coincide with phylogenetic affiliation.
Phylogeny and siderophore production of Cytophaga isolates
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Siderophore production capabilities, type of siderophore produced, depth of isolation, and
station are also indicated. Isolates are not as closely related as in other groups such as
Pseudoalteromonasand Alteromonas. This phylogenetic group only appeared below the
chlorophyll ihaximum.
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Discussion:
This library of isolates presented a unique opportunity for exploring the heterogeneity of
iron scavenging systems within easily cultured bacterial populations in this region. Based
on fluorescence measurements, Stations 5 and 7 appeared to be the most enveloped by
the CRD. They also had the shallowest chlorophyll maxima at 35 m and 34 m,
respectively. Stations 2 and 15, while still in a highly productive area, are on the outskirts
of the CRD. They had slightly deeper chlorophyll maxima at 51 m and 46 m. Station 17
was well outside the CRD in a predicted HNLC region, and had the deepest chlorophyll
maximum at 55m. The water column structure shifts between these stations suggests that
the samples did in fact move through the upwelling and into nutrient depleted waters.
These spatial and structural differences may well account for the shifts in the relative
abundance of siderophore producing isolates between stations. Station 2, for example, has
no siderophore production by isolates in shallow 8m and 35m waters. The depth of peak
siderophore producing capabilities in the isolate library shifted deeper as the ship traveled
through the CRD, and shoaled again as the water became more nutrient-depleted.
While it is possible to use the CAS assay to quantify siderophores when used with pure
isolations, quantification in environmental samples with unknown ligand composition has
yet to be evaluated for reliability. The CAS assay indicates a compound's ability to
remove iron loosely bound to the dye complex, and so can indicate the presence of both
siderophore and other weaker iron-binding compounds. Uninoculated media,
uninoculated media containing dipyridyl, and desferral were used as two negative and
one positive control to rule out non-bacterial compounds as false positives. Czsaky and
Arnow assays carried out on sequenced isolates suggest that there was siderophore
production in the isolates, and that the CAS results were not simply other weaker ironbinding molecules. The fact that the results of the CAS assays were repeated with
accuracy suggests that this is not an artifact of variable experimental conditions.
All stations contained at least one depth where the isolates were overwhelmingly
siderophore-producers, suggesting a unique competitive structure exists at these depths.
An increase in siderophore producing isolates could indicate that iron acquisition in that
zone of the water column is more competitive, or that biologically-produced ligands are
more successful in making iron bioavailable than in other zones in representatives from
this library. Data on cobalt-binding and iron-binding ligands from this region suggests
that there are trace-metal binding ligands in surface waters, which shows that while this
library provides a window to microbial siderophore production in the ocean, it is not a
complete picture (Saito et al. 2005, Rue unpublished).
Closely related isolates within the same group have different siderophore-producing
capabilities, which suggests that there is a lot of genetic heterogeneity for this metabolic
pathway. While the cellular biomass of the cultures tested for siderophore production was
not normalized, all culture wells showed turbidity and 99.2% of reactions were decisive
enough to be counted. The data shown in Figure 13 indicates that there are sequences
from isolates within Alteromonas that are very closely related yet have differing
siderophore producing capabilities. This trend is carried throughout the different
phylogenetic groups represented by isolates in this study, suggesting genetic microheterogeneity.
Another interesting result is that under our experimental conditions, isolates of known
siderophore producers did not always produce siderophores. Halomonas,Alteromonas,
and Marinobacterisolates are representatives in this category. It is plausible - likely,
even - that since our knowledge of these microbes' siderophore producing capabilities is
based on a limited number of laboratory species and strains, that the range of their
metabolic capabilities in the field is not fully characterized. Different strains may have
different iron stress thresholds for siderophore gene expression. Furthermore, a negative
result from a CAS assay does not rule out siderophore production - it may be that the
siderophores are too dilute, are too weak to cause a color change, or react with iron in a
way the assay cannot detect. The data certainly indicates that individual strains have the
ability to compete for iron in different ways, even while using the same general strategy.
This speaks to the complexity of biological impact on nutrient cycling in general - and
the difficulty in teasing the microbial loop apart to find a clearer picture.
Just as with overall siderophore production patterns versus phylogeny, the type of
siderophore produced seemed to be heterogeneous within taxonomic groups. When
looking at the phylogenetic trees (Figures 13-19), Marinobacter(Figure 15) and
Idiomarina (Figure 16) were the only taxonomic groups that did not include both
catechol and hydroxamate-type siderophore producing strains under our experimental
conditions. Within each group that contained both siderophore types, Pseudoalteromonas
is the only one that seems to have a distinct grouping of catechol vs hydroxamate
producers. This general lack of phylogenetic grouping suggests a high diversity of
siderophore producing capabilities, and a lack of competitive advantage of one structure
versus another. It also suggests a high rate of transfer of genes related to siderophore
production within and between these groups, but more information is needed to confirm
this.
High numbers of catechol producers in surface waters is unexpected due to the theory
that catechol-type siderophores are more susceptible to photooxidation, and thus are
predicted to be less prevalent in the photic zone than hydroxamate siderophores (Barbeau
et al. 2003). The trend found in this study may be a result of the culturing bias of the
isolate library obtained in this study. Because all siderophore producing microbes are not
accounted for in this library, it is impossible to predict what trends would appear if the
entire siderophore pool had been analyzed at each depth and station. Quantitative
analysis to determine how large a fraction of the microbial population the isolates made
up would provide insight into their influence on environmental siderophore production.
Since uptake and utilization strategies have not been well characterized in marine
bacteria, it is not yet possible to use these chemical properties for predicting how the
photolabile nature of certain siderophores influences biological production. It is possible
that the high turnover of these compounds could render their susceptibility to
photodegredation unimportant from a biological standpoint. Studies on how structural
changes affect siderophore receptor recognition would be the first step in understanding
the link between chemical properties and biological function.
Because the Alteromonas and Pseudoalteromonaswere the most highly represented
groups, distinct hydroxamate and catechol production patterns were observed in the depth
profile. Alteromonas was dominated by hydroxamate production, especially at depth.
Catechol production only occurred in the shallowest three depths. Pseudoalteromonas
was dominated by catechol production, with hydroxamates only being produced at 35m
and 150m. Although these groups are very closely related, they had different dominating
siderophore structural classes. Again, the groups showed heterogeneity in their structural
production. More sequenced representatives from the other groups would be an excellent
way to compare their siderophore structural production patterns.
RFLP patterns of 16S rRNA genes produced by this isolate library also show
phylogenetic heterogeneity. While each group had different proportions of isolates within
the different patterns, no group had less than three patterns represented. As Figures 13-17
show, the largest groups of isolates had members distributed widely across the patterns.
This genetic diversity is also clear when the patterns of catechol versus hydroxamate
producers are examined. Although each has three pattern groups exclusive to itself, they
both have similar proportions among their shared pattern groups. A more detailed RFLP
analysis at higher resolution could provide more information on subtle differences that
occur among and between these groups.
By characterizing the siderophores produced by these isolates and determining the
genetic make-up of the population, we can better understand the impact of siderophore
production on iron cycling in this region. This appears to be the first large-scale genetic
and siderophore production assessment of a heterotrophic population within both an
upwelling and predicted high nutrient low chlorophyll regime (HNLC). One of the
exciting aspects of marine microbiology is that many environmental processes are still, in
a broader sense, not yet well characterized. Isolating specific components of a microbial
community provides is a unique opportunity to manipulate, stress, and test their
metabolic capabilities. The data derived from isolate collections such as this represents a
step towards a better understanding of organisms and processes in the field. This sets the
stage to study actual siderophore production - or other metabolic processes - in situ. By
combining cultivation techniques with modem molecular methods, a more thorough
understanding of bacterial processes in the environment, and their contribution to
community interactions and biogeochemical processes is now becoming more possible.
The library of isolates described in this study should continue to provide opportunities to
gain further insight into siderophore occurrence, distribution, and diversity. More
complete structural analysis is possible with these laboratory models, which would add a
substantial amount to current understanding of siderophore structure. These isolates may
also facilitate gene knockout experiments, which could provide new gene markers to
assess siderophore production, and possibly iron stress in the field. In summary, the work
described in this study should facilitate new approaches to monitoring siderophore
production in the field and better understand how they contribute to microbial survival,
competition, and oceanic iron cycling.
Acknowledgements:
I would like to thank Eric Webb for his leadership and support during this project. Adam
Rivers, Dreux Chappell, Annette Hynes, and Nan Trowbridge were invaluable lab
members. I would also like to thank Mak Saito, the chief scientist on the cruise for this
project, as well as his laboratory for continued support. I thank John Waterbury for
providing lab space as well as for being my advocate. I would like to thank Ed DeLong
and Tracy Mincer for their help with the phylogenetic analysis in this project. I express
thanks to the Captain and crew of the R/V Knorr for an exceptional cruise. I am grateful
for the use of Dave Caron and Karen Casiotti's gel imaging system and gel box. I also
thank the Academic Programs staff and my fellow Biology third years for their guidance
and support.
I am grateful to my funding sources, National Science Foundation grants BO OCE0352241 "The Effect of Iron Bioavailability on Synechococcus diversity from a HNLC
region to the Costa Rica upwelling dome" and CO-OCE-0452883 "Interactions of Cobalt
and Iron with in situ Cyanobacterial Physiology in the South Atlantic and the Benguela
Upwelling Region", the National Science Foundation Graduate Research Fellowship, and
the Academic Programs office.
Works Cited:
Andrews, S.C., Robinson, A.K. and F. Rodriquez-Quinones, 2003. Bacterial iron
homeostasis. FEMS Microbiol Rev, 27: 215-237.
Armstrong, J.E., and C. Van Baalen, 1979. Iron transport in microalgae: the isolation and
biological activity of a hydroxamate siderophore from the blue-green alga
Agmenellum quadruplicatum.J. Gen. Microbiol. 111, 253-262.
Arnow, E., 1937. Colorimetric determination of the components of 3, 4Dihydroxyphenylalaninetyrosine mixtures. J. Biolog. Chem., 531-537.
Barbeau, K., E. L. Rue, C. G. Trick, K. W. Bruland, and A. Butler, 2003. Photochemical
Reactivity of Siderophores Produced by Marine Heterotrophic Bacteria and
Cyanobacteria Based on Characteristic Fe(III) Binding Groups. Limnol.
Oceanogr. 48: 1069-1078.
Barber, R.T., and J.H. Ryther, 1969. Organic chelators: Factors affecting primary
production in the Cromwell Current upwelling. J. Exp. Mar. Biol., 3: 191-199.
Beja, O., Spudich, E.N., Spudich, J.L., Leclerc, M., and E.F. DeLong, 2001.
Proteorhodopsin phototrophy in the ocean. Nature, 411: 786-789.
Bickel, H., Hall, G.E., Keller-Schierlein, W., Prelog, V., Visher, E. and A. Wettstein,
1960. Stoffwechselprodukte von actinomyceten. Uber die konstitution von f
errioxamin B. Helv. Chim. Acta, 43: 2129-2138.
Braun, V., Hantke, K. and W. Koster, 1998. Bacterial iron transport: Mechanisms,
genetics, and regulation. In A. Siegel and H. Sigel (eds.), Iron transport and
storage in microorganisms, plants and animals, v. 35. Marcel Dekker, pp. 67-145.
Braun, V. and H. Killman, 1999. Bacterial solutions to the iron-supply problem. Trends
Biochem. Sci., 24:3389-3402.
Broenkow, W.H., 1965. The distribution of nutrients in the Costa Rica Dome in the
Eastern Tropical Pacific Ocean. Limnol. Oceanogr., 10(1): 40-52.
Bruland, K.W., Orians, K.J., J.P. Cowen, 1994. Reactive trace metals in the stratified
central North Pacific. Geochim. Cosmochim. Acta, 58: 3171-3182.
Buchanan, S.K. and others, 1999. Crystal structure of the outer membrane active
transporter FepA from Esherichia coli. Nat. Struct. Biol., 6:56-63.
Campbell, L., Nolla, H.A., and D. Vaulot, 1994. The importance of Prochlorococcusto
community structure in the central North Pacific Ocean. Limnol. Oceanogr., 39:
954-961.
Coale, K.H., et al., 1996. A massive phytoplankton bloom induced by an ecosystem-scale
iron fertilization experiment in the equatorial Pacific Ocean. Nature, 383: 495501.
Crichton, R.R., and R.J. Ward, 1992. Iton metabolism - new perspectives in view.
Biochemistry, 31: 11255-11264.
Crichton, R.R., and R.J. Ward, 1998. Iron homeostasis. Met Ions Biol Syst, 35: 633-635.
Csaky, T.Z., 1948. On the estimation of bound hydroxylamine in biological materials.
Acta Chem. Scand., 2: 450-454.
de Baar, H.J.W., Buma, A.G.J., Nolting, R.F., Cadee, G.C., Jaques, G. and P.J. Treguer,
1990. On iron limitation of the Southern Ocean: Experimental opservations in the
Weddell and Scotia Seas. Mar. Ecol. Prog. Ser., 65: 105-122.
DuRand, M.D., Olson, R.J. and S.W. Chisholm, 2001. Phytoplankton population
dynamics at the Bermuda Atlantic time series station in the Sargasso Sea. DeepSea Res. II, 48: 1983-2003.
Ehrenreich, I.M., Waterbury, J.B. and E.A. Webb, 2005. Distribution and diversity of
natural product genes in marine and freshwater cyanobacterial cultures and
genomes. Appl. Environ. Micro., 71(11): 7401-7413.
Ferguson, A.D., Hofmann, E., Coulton, J.W., Diederichs, K. and W. Welte, 1998.
Siderophore-mediated iron transport: Crystal structure of FhuA with bound
lipopolysaccharide. Science, 282: 2215-2220.
Fredrickson, J., Zachara, J., Balkwill, D. et al., 2004. Geomicrobiology of high-level
nuclear waste-contaminated vadose sediments at the Hanford site, Washington
state. Appl. Environ. Microbiol., 70 (7): 4230-41.
Gledhill, M. and C.M.G. van den Berg, 1994. Determination of complexation of iron(III)
with natural organic complexing ligands in sea water using cathodic stripping
voltammetry. Mar. Chem., 47: 41-54.
Goericke, R. and N. Welschmeyer, 1993. The marine prochlorophyte Prochlorococcus
contributes significantly to phytoplankton biomass and primary production in the
Sargasso Sea. Deep-sea Res. I, 40: 2283-2294.
Gordon, R.M., Martin, J.H., G.A. Knauer, 1982. Iron in north-east Pacific waters. Nature,
299: 611-612.
Granger, J., and N. M. Price. 1999. The Importance of Siderophores in Iron Nutrition of
Heterotrophic Marine Bacteria. Limnol. Oceanogr. 44:541-555.
Haygood, M.G., Holt, P.D., and A. Butler, 1993. Aerobactin production by a planktonic
marine Vibria sp., Limnol. Oceanogr., 38(5): 1091-1097.
Hider, R.C., 1984. Siderophore mediated absorption of iron. Struct. Bond. Berlin, 58: 2687.
Hofmann, E.E., Busalacchi, A.J. and J.J. O'Brien. Wind Generation of the Costa Rica
Dome. Science, 214: 552-554.
Hudson, R.J.M., 1998. Which aqueous species control the rates of trace metal uptake by
aquatic biota? Observations and predictions of non-equilibrium effects. Sci. Total
Environ., 219: 95-115.
Hutchins, D.A., Witter, A.E., Butler, and G.W. Luther III, 1999. Competition among
marine phytoplankton for different chelated iron species. Nature 400, 858-861.
Kirchman, D.L., Meon, B., Cottrell, M.T., Hutchins, D.A., Weeks, D., K.W. Bruland,
2000. Carbon versus iron limitation of bacterial growth in the Californian
upwelling regime. Limnol. Oceanogr., 45: 1681-1688.
Landing, W.M., and K.W. Bruland, 1987. The contrasting biogeochemistry of iron and
manganese in the Pacific Ocean. Geochim. Cosmochim. Acta, 51: 29-43.
Lewis, B.L., P.D. Holt, S.W. Taylor, S.W. Wilhelm, C.G. Trick, A. Butler, and G.W.
Luther III. Voltammetric etimation of Fe(III) thermodynamic stability constants
for catecholate siderophores isolated from marine bacteria and cyanobacteria.
Marine Chemistry 50 (1995), 179-188.
Lewis, B.L., Luther III, G.W., Lane, H. and T.M. Church, 1995. Determination of metalorganic complexation in natural waters by SWASV with pseudopolarograms.
Electroalalysis, 7: 166-177.
Lewis, B.L., Holt, P.D., Taylor, S.W., Wilhelm, S.W., Trick, C.G., Butler, A. and G.W.
Luther III, 1995. Voltammetric estimation of iron(III) thermodynamic stability
constants for catecholate siderophores isolated from marine bacteria and
cyanobacteria. Mar. Chem., 50: 179-188.
Macrellis, H.M., Trick, C.G., Rue, E.L., Smith, G., and K.W. Bruland, 2001. Collection
and detection of natural iron-binding ligands from seawater. Marine Chem., 76:
175-187.
Madigan, M. T., 2003. Anoxygenic phototrophic bacteria from extreme environments.
Photosynthesis Research, 76: 157-171.
Maldonado, M.T., and N.M. Price, 1999. Utilization of iron bound to strong organic
ligands by plankton communities in the subarctic Pacific Ocean. Deep-Sea Res. !,
46: 2447-2473.
Martin, J.H. and S.E. Fitzwater, 1988. Iron deficiency limits phytoplankton growth in the
north-east Pacific subarctic. Nature, 331: 341-343.
Martin, J.H., Gordon, R.M., Fitzwater, S., W.W. Broenkow, 1989. VERTEX:
phytoplankton/iron studies in the Gulf of Alaska. Deep-Sea Res. 36, 649-680.
Martin, J.H., Gordon, R.M., S.E. Fitzwater, 1991. The case for iron. Limnol. Oceanogr.,
36: 1793-1802.
Martin, J.H. et al., 1994. Testing the iron hypothesis in ecosystems of the equatorial
Pacific Ocean. Nature, 371: 123-129.
Martinez, J.S., Zhang, G.P., Holt, P.D., Jung, H.-T., Carrano, C. J., Haygood, M. G. and
A. Butler, 2000. Self-assembling amphiphilic siderophores from marine bacteria.
Science, 287: 1245-1247.
Martinez, J. S., M. G. Haygood, and A. Butler. 2001. Identification of a natural
desferrioxamine siderophore produced by a marine bacterium. Limnol. Oceanogr.
Martinez, J. S., J. N. Carter-Franklin, E. L. Mann, J. D. Martin, M. G. Haygood, and A.
Butler. 2003. Structure and Membrane Affinity of a Suite of Amphiphilic
Siderophores Produced by a Marine Bacterium. Proceedings of the National
Academy of Sciences of the United States of America 100: 3754-3759.
Moore, J. K., Doney, S. C. and K. Lindsey, 2004. Upper ocean ecosystem dynamics and
iron cycling in a global three-dimensional model. Global Biogeochem. Cycles, 18:
GB4028.
Morel, F.M.M., N.M. Price, 2003. The biogeochemical cycles of trace metals in the
oceans. Science, 300: 944-947.
Neilands, J.B., 1984. Methodology of siderophores. Struct. Bond. Berlin, 58: 1-24.
Neilands, J.B., and K. Nakamura, 1991. Detection, determination, isolation,
characterization and regulation of microbial iron chelates. In: WInkelman, G.
(Ed.), CRC Handbook of Microbial Iron Chelates. CRC Press, Boca Raton, FL,
pp. 1-14.
O'Brien, I.G. and F. Gibson, 1970.The structure of enterochelin and related 2,3dihydroxy-N-benzoylserine conjugates from Escherichiacoli. Biochem. Biophys.
Acta, 215: 393-402.
Pollack, J.R. and J.B. Neilands, 1970. Enterobactin, an iron transport compound from
Salmonella typhimurium. Biochem. Biophys. Res. Commun., 38: 989-992.
Ratledge, C. and L.G. Dover, 2000. Annu. Rev. Microbiol., 54: 881-941.
Rue, E.L., and K.W. Bruland, 1995. Complexation of iron(III) by natural organic ligands
in the Central North Pacific as determined by a new competitive ligand
equilibration/adsorptive cathodic stripping voltammetric method. Mar. Chem., 50:
117-138.
Rue, E.L., and K.W. Bruland, 1997. The role of organic complexation on ambient iron
chemistry in the equatorial Pacific Ocean and the response of a mesoscale iron
addition experiment. Limnol. Oceanogr., 42: 901-910.
Ryther, J.H., and D.D. Kramer, 1961. Relative iron requirement of some coastal and offshore plankton algae. Ecology, 42: 444-446.
Saito, M.A., Rocap, G. and J.W. Moffett, 2005. Production of cobalt binding ligands in a
Synechococcus feature at the Costa Rica upwelling dome. Limnol. Oceanogr.,
50(1): 279-290.
Schwyn, B., and J.B. Neilands, 1987. "Universal CAS assay for the detection and
determination of siderophores". Anal. Biochem., 160: 47-60.
Sugiuira Y. and K. Nomoto, 1984. Phytosiderophores: Structures and properties of
mugineic acids and their metal complexes. Struct. Bond. Berlin, 58: 107-135.
Sunda, W.G. and S.A. Huntsman, 1995. Cobalt and zinc interreplacement in marine
phytoplankton: Biological and geochemical implications. Limnol. Oceanogr., 40:
1404-1417.
Tortell, P.D., M.T. Maldonado, and N.M. Price. 1996. The role of heterotrophic bacteria
in iron-limited ocean ecosystems. Nature 383: 330-332.
van der Helm, D., 1998. The physical chemistry of bacterial outer-membrane siderophore
receptor proteins. In A. Siegl and H. Sigel (eds.), Iron transport and storage in
microorganisms, plants, and animals, pp. 355-401.
Wagner-Dobler, I, H Rheims, D A Felske, R Pukall and B J Tindall, 2003. "Jannaschia
helgolandensis gen. nov., sp. nov., a novel abundant member of the marine
Roseobacter clade from the North Sea". Int. J. System.
Evol. Micro., 53: 731-738.
Whitman, W., Coleman, D., and W. Wiebe, 1998. Prokaryotes: The unseen majority.
Proc. Natl. Acad. Sci., 95 (12): 6578-6583.
Wilhelm, S.W., and C.G. Trick, 1994. Iron-limited growth of cyanobacteria: multiple
siderophore production is a common response. Limnol. Oceanogr. 39, 1979-1984.
Wilhelm, S.W. and C.G. Trick, 1995. Physiological profiles of Synechococcus
(Cyanophyceae) grown in iron-limited continuous culture. J. Phycol., 31(1): 7985.
Wilhelm, S.W., MacAuley, K., and C.G. Trick, 1998. Evidence for the importance of
catechol-type siderophores in the iron-limited growth of a cyanobacterium.
Limnol. Oceanogr., 43: 992-997.
Wu, J., and G.W. Luther, 1995. Complexation of Fe(III) by natural organic ligands in the
northwest Atlantic Ocean by a competitive ligand equilibrium method and a
kinetic approach, Mar. Chem., 50: 159-177.