Geobiology (2011), 9, 360–375
DOI: 10.1111/j.1472-4669.2011.00283.x
Low temperature S0 biomineralization at a supraglacial spring
system in the Canadian High Arctic
D. F. GLEESON,1 C. WILLIAMSON,2 S. E. GRASBY,3 R. T. PAPPALARDO,1 J. R. SPEAR2 AND
A. S. TEMPLETON3,4
1
Jet Propulsion Laboratory, California Institute of Technology, Pasadena, California, USA
Colorado School of Mines, Golden, Colorado, USA
3
Geological Survey of Canada, Calgary, Alberta, Canada
4
University of Colorado, Boulder, Colorado, USA
2
ABSTRACT
Elemental sulfur (S0) is deposited each summer onto surface ice at Borup Fiord pass on Ellesmere Island, Canada,
when high concentrations of aqueous H2S are discharged from a supraglacial spring system. 16S rRNA gene
clone libraries generated from sulfur deposits were dominated by b-Proteobacteria, particularly Ralstonia sp. Sulfur-cycling micro-organisms such as Thiomicrospira sp., and e-Proteobacteria such as Sulfuricurvales and Sulfurovumales spp. were also abundant. Concurrent cultivation experiments isolated psychrophilic, sulfide-oxidizing
consortia, which produce S0 in opposing gradients of Na2S and oxygen. 16S rRNA gene analyses of sulfur precipitated in gradient tubes show stable sulfur-biomineralizing consortia dominated by Marinobacter sp. in association with Shewanella, Loktanella, Rubrobacter, Flavobacterium, and Sphingomonas spp. Organisms closely
related to cultivars appear in environmental 16S rRNA clone libraries; none currently known to oxidize sulfide.
Once consortia were simplified to Marinobacter and Flavobacteria spp. through dilution-to-extinction and agar
removal, sulfur biomineralization continued. Shewanella, Loktanella, Sphingomonas, and Devosia spp. were
also isolated on heterotrophic media, but none produced S0 alone when reintroduced to Na2S gradient tubes.
Tubes inoculated with a Marinobacter and Shewanella spp. co-culture did show sulfur biomineralization, suggesting that Marinobacter may be the key sulfide oxidizer in laboratory experiments. Light, florescence and scanning electron microscopy of mineral aggregates produced in Marinobacter experiments revealed abundant cells,
with filaments and sheaths variably mineralized with extracellular submicron sulfur grains; similar biomineralization was not observed in abiotic controls. Detailed characterization of mineral products associated with low temperature microbial sulfur-cycling may provide biosignatures relevant to future exploration of Europa and Mars.
Received 10 May 2010; accepted 03 April 2011
Corresponding author: D. F. Gleeson. Tel.: 818 354 2786; fax: 818 354 2494; e-mail: Damhnait.F.Gleeson@
jpl.nasa.gov
INTRODUCTION
While uncommon on Earth at the present day, cold environments dominated by sulfur chemistry could form critical
habitats for potential life at other locations in the solar system,
such as Europa and Mars (Kargel et al., 2000; Zolotov &
Shock, 2004; Langevin et al., 2005). At the surface of
Europa, probable sulfur-rich, non-ice materials are concentrated along geologic features (McCord et al., 1998; Carlson
et al., 1999; Dalton, 2007) and may reflect the chemistry of
an ocean in communication with the surface. Materials carried
to the surface by partial melt or mobile ice from deeper ice or
ocean environments could be investigated for the presence of
360
biosignatures by future missions (Dalton et al., 2003; ChelaFlores, 2006). H2S and elemental sulfur (S0) may have
provided important energy sources for early metabolisms on
the Earth once strong oxidants such as O2 and NO3) became
available after the great oxidation event more than two billion
years ago. H2S accumulation and persistence in euxinic deep
ocean waters may have extended into the Neoproterozoic
(1.0–0.54 Gyr), as proposed by Canfield (1998). Evidence
for global glaciations on the Earth in the late Neoproterozoic
(Hoffman et al., 1998; Hoffman & Schrag, 2002) indicate
that low temperature conditions extended over the majority
of the Earth’s surface and psychrophilic sulfide oxidation may
have gained increased importance under such conditions.
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Low temperature S0 biomineralization
Despite their geobiological and astrobiological importance,
micro-organisms that can conserve chemical energy from oxidation of reduced sulfur compounds under psychrophilic conditions have been little studied to date. In addition, the
activities of sulfur-generating microbial communities that may
produce microscopic and macroscopic mineral deposits with
detectable biosignatures has until recently remained relatively
unexplored (Gleeson et al., in revision). Large-scale mineral
deposits can be remotely observed on Earth and other planetary bodies (Kruse et al., 1990; Mccord et al., 1999; Gendrin
et al., 2005; Gleeson et al., 2010), and could potentially be
utilized to provide targets for in situ investigation of signs of
microbial activity (Figueredo et al., 2003; Chela-Flores,
2006). The targets could include micro-fabrics characteristic
of microbialites (Grotzinger & Knoll, 1999), or isotopic signatures, chemical compositions or mineralogical structures
indicative of biomineralization (Banfield et al., 2001). The
primary challenge inherent in the use of S0 as a biosignature is
the lack of characterization of the micro-fabrics or morphologies associated with microbial S0 formation, as well as an
understanding of the environmental and biological conditions
under which S0 is stabilized and preserved.
Geologic deposits of S0 form under a wide range of environmental conditions, such as in molten volcanic flows
(Wantanabe, 1940), at fumaroles (Ljunggren, 1960), at
hydrothermal vents (Taylor et al., 1999), within marine sediments (Thamdrup et al., 1994), and over salt domes and
evaporite deposits (Davis & Kirkland, 1979), with the potential for many of these to be microbially mediated. The 8-electron reduction of sulfate to sulfide does not result in the
formation of S0 as an intermediate (Troelsen & Jorgensen,
1982; Machel, 2001); instead, S0 typically indicates the
incomplete microbial or abiotic oxidation of reduced sulfur
species such as H2S. The complete oxidation of H2S to SO42)
occurs in a series of steps, frequently forming S0 as an intermediate (Fuseler & Cypionka, 1995; Ehrlich, 2002).
Microbiological studies of warm sulfide springs associated
with magmatic activity in locations such as Yellowstone
National Park, Wyoming (Farmer & Des Marais, 1994b; Fouke
et al., 2001), and Jasper National Park in Alberta, Canada
(Grasby et al., 2000; Bonny & Jones, 2003) have explored the
role that microbes may play in the precipitation of sulfur and
carbonate minerals, while cold sulfide spring sites remain relatively understudied to date. Prior to the use of 16S rRNA technology, Thermothrix within the b-Proteobacteria were thought
to be the dominant sulfur oxidizers at hot spring sites such as
Mammoth in Yellowstone although clone libraries of DNA
extracted from sulfur streamers at such sites were subsequently
shown to be dominated by Aquificales while b-Proteobacteria
were absent (Brenner et al., 2005). Reed et al. (2006) explored
the microbial diversity of Bacteria and Archaea in sulfide coldseep sediments at the base of the Florida Escarpment. Douglas
& Douglas (2000) used light and electron microscopy to visually explore the microbiology and mineralogy of a cold (9 "C)
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361
anoxic sulfur spring in Ancaster, Ontario, Canada. In a stream
channel, they observed colloidal sulfur-associated with unicellular, colorless, sulfur-oxidizing bacteria potentially responsible for the production of macroscopic filamentous structures.
Perreault et al. (2007, 2008) found that in clone libraries constructed from cold sulfide springs emerging from permafrost
on Axel Heiberg Island, Nunavut, 16S rRNA sequences from
the bacterial community group with known autotrophic and
heterotrophic sulfide and sulfur oxidizers. In their study, Thiomicrospira arctica, Thiobacillus sp. strain EBD bloom, and
Halothiobacillus sp. strain RA13, accounted for 30–74% of the
microbiological community. Neiderberger et al. (2009) also
isolated several autotrophic sulfide and thiosulfate oxidizing
Thiomicrospira sp. from samples of microbial streamers collected at the same site.
The supraglacial spring system of Borup Fiord Pass
at 81"N, 81"W on Ellesmere Island in the Canadian High
Arctic (Grasby et al., 2003) is a unique low temperature
environment where S0 is found in abundance (Fig. 1).
Subsurface saline fluids rise through an estimated 200 m-thick
glacier and deliver sulfur-rich waters that have mixed with glacial melt water, and deposit S0, gypsum (CaSO4.2H2O) and
calcite (CaCO3) on the surface as well as along flow channels
incised into the ice (Fig. 1C). The unstable CaCO3 polymorph vaterite has also been observed (Grasby, 2003). Sulfide
and sulfate levels within the springs have been measured as
high as 142 mg L)1 (4.2 mM) and 1786 mg L)1 (27.9 mM),
respectively, which are the highest measured dissolved sulfur
concentrations across the Canadian Arctic Islands. Measured
temperatures for the springs are !0 "C while the fluid pH is
alkaline between 8 and 9. The water pH rapidly decreases to
!pH 6 and H2S(aq) drops below detection within meters of
the spring source. Sulfur isotopic evidence suggests that the
source of sulfur is most likely sedimentary sulfate (anhydrate,
gypsum) that lies at depth beneath the base of the glacier
(Grasby et al., 2003).
Microbial organisms have previously been detected within
the spring waters and precipitates in cell densities of 1.9–
2.9 · 104 cells mL)1 (Grasby et al., 2003). Limited investigations of microbiological community composition within
the precipitates have revealed the presence of c-Proteobacteria
closely related to Marinobacter sp. (98% identity match) by
cloning and 16S rRNA sequencing of extracted DNA, while
sequencing of bands observed in denaturing gradient gel electrophoresis (DGGE) found close matches with the b-Proteobacteria Polaromonas (99% match), Burkholderia sp., and the
c-Proteobacterium Pseudomonas fulva (Grasby et al., 2003).
Due to the limited microbial community data for the site and
the dynamic nature of the spring system, little is known about
the microbial ecology at the spring.
The environmental and microbiological factors that lead to
the formation of S0 on the surface of the glacier at Borup
Fiord Pass have not been previously investigated in detail. Our
objective was to explore the microbial diversity of sulfur-
362
D. F . GLEE SON e t al.
A
C
associated microbial communities, and to conduct targeted
cultivation of psychrophilic micro-organisms that play a key
role in the production of oxidized, sulfur-rich deposits on
the surface ice. In particular, by combining microbiological,
geochemical and mineralogical data, we expand our knowledge of biologically mediated sulfur-cycling processes in cold
environments, which could inform the search for S-based
microbial life and biosignatures in other locations in the solar
system.
MATERIALS AND METHODS
Geochemical and mineralogical analysis of site material
Water samples and moist mineral precipitates along channels
were collected from the actively flowing spring site and associated sulfur deposits during a field expedition to Borup Fiord
Pass in June and July of 2006. Unstable parameters (temperature, pH, dissolved HS)) were measured at sites along the
B
Fig. 1 Borup Fiord pass, Ellesmere Island. (A)
Landsat imagery of the N-S trending valley and central glaciers in 2005. (B) Local geology of the same
area from Gleeson et al. (2010), modified from
Thorsteinsson (1974). Springs and associated sulfur
deposits on the glacier are indicated in red. (C) View
down-glacier of deposits of elemental sulfur,
gypsum and calcite on glacial ice (up-glacier spring
source not shown). Image obtained June 2006.
channel, and at increasing distances from the discharge site
(Fig. 2). Dissolved HS) was measured using CHEMetrics#
(Calverton, VA, USA) colorimetric kits (±0.01 mg L)1). For
chemical analyses, water samples were passed through
0.45-lm filters and stored in the dark at 4 "C in high-density
polyethylene bottles until analyzed. Samples for cation
analyses were acidified with ultrapure nitric acid to pH <2.
Subsequent chemical analyses were carried out at the
Geological Survey of Canada. Alkalinity was determined by a
standard end-point titration. Anions were measured by ion
liquid chromatography, and cations were measured by inductively coupled plasma emission spectrometry. Analytical error
was estimated to be less than 2% (Table 1).
Mineralogy of precipitates collected from the deposits was
determined by X-ray diffraction (XRD) analysis on a Philips
PW1700 powder diffraction system with Cobalt x-ray source.
All analyses were on powder samples, which were executed
by the PANalytical X’Pert Quantify software; samples
were usually scanned from 2–60" 2Ø at 1" per minute with
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Low temperature S0 biomineralization
363
Table 2 X-ray diffraction mineralogical measurements on field samples from
2006
ID
Lat
Long
Gypsum
Sulfur
Quartz
Calcite
BF06-04A
BF06-05
BF06-05B
81"01.712¢
81"01.615¢
81"01.615¢
81"37.412¢
81"36.708¢
81"36.708¢
<1
<1
<1
45
16
48
trace
10
<1
55
74
52
Numbers represent percentages of total, the locations of which are shown in
Fig. 2.
Fig. 2 Sample locations at increasing distance from the spring source, collected
at sites of deep (a few cm thick) mineral deposits.
Table 1 2006 spring water geochemistry
Ion
mg L)1
Alk*
F)
Cl)
Br)
NO3)
PO42)
SO42)
Na+
Ca2+
Mg2+
K+
TDS!
57.8
2.97
3234
0.00
0.00
0.00
1786
1248
696
353
16.26
7394
*Alk, alkalinity; !TDS, total dissolved solids.
accelerating voltage of 40 KV and 30 mA. Mineral determination was processed by PANalytical’s X’pert Highscore program, and the quantification of mineral composition within
each sample was calculated from their mineral peak intensities
(or peak area). The whole rock results are semi-quantitative
and they are expressed in mineral ratio percent (Table 2).
Environmental DNA clone libraries
DNA was extracted from !0.5 g of mineral samples 4A and
5B (locations shown in Fig. 2) that were preserved in 70%
ethanol using a phenol chloroform extraction technique.
Polymerase chain reaction (PCR) was then conducted using
protocols described by Sahl et al. (2010). The PCR
primers 515F (5¢-GTGCCAGCMGCCGCGGTAA-3¢) and
1391R (5¢-GACGGGCGGTGWGTRCA-3¢) were selected
to amplify small-subunit rRNA gene fragments for all three
domains of life (Lane, 1991). After this initial screen of diversity, the Bacteria-specific forward primer 8F (5¢-AGAGTTTGATCCTGGCTCAG-3¢) was used in conjunction with the
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universal reverse primer 1492R (5¢-GGTTACCTTGTTACGACTT-3¢) to obtain full-length bacterial 16S rRNA gene
sequences (Lane, 1991); while PCR conducted with archaealspecific primers was unable to amplify small-subunit rRNA
gene fragments. Appropriate negative controls were employed
throughout the DNA extraction and PCR processes. PCR
products were gel purified with the Montage DNA Gel
Extraction Kit (Millipore, Billerica, MA, USA), cloned with
the TOPO TA Cloning Kit (Invitrogen, Carlsbad, CA, USA)
using electrocompetent Escherichia coli TOP10 cells, and
sequenced on a MegaBACE 1000 dye-terminating sequencer.
Sequences were assembled with Xplorseq (Frank, 2008), and
screened for chimeras with Mallard (Ashelford et al., 2006).
The resulting 349 non-chimeric sequences were aligned with
the NAST aligner (Desantis et al., 2006b), and these alignments were manually checked in ARB (Ludwig et al., 2004).
Phylogenetic relationships of sequences were determined by
inserting the sequences into the ARB dendrogram by parsimony insertion using the Lanemask filter (Lane, 1991) and
the most recent Greengenes (Desantis et al., 2006a) database
release. Relevant sequences were exported from ARB with the
Lanemask filter, and phylogenetic trees were constructed with
Mr Bayes (Huelsenbeck et al., 2001).
Targeted culture of sulfide-oxidizers that produce S0
Sample material collected aseptically with alcohol and flame
sterilized spatulas from spring deposits in the 2006 field season
was used to inoculate sulfide gradient tubes on return to the
laboratory. Gradient tubes were assembled using iron sulfide
plugs created by adding FeS precipitate prepared according to
the method of (Hanert, 2006) in a 1:1 ratio with EM media
(see below) and high melt agarose (1% wt ⁄ vol) in a flask. 1 mL
aliquots of the autoclaved plug material were allowed to cool
in 10 mL glass tubes and covered by 5 mL of a semi-solid agar
slush overlayer (0.15 wt ⁄ vol agarose) of an autoclaved, artificial seawater medium (EM) containing 27.5 g NaCl, 5.38 g
MgCl2, 0.72 g KCl, 0.2 g NaHCO3, 1.4 g CaCl2, 1 g
NH4Cl, and 0.05 g K2HPO4, dissolved above in 967 mL
deionized H2O. The medium was amended with 1 mM
NaHCO3, and sterile N2:CO2 was also bubbled through the
agar overlayer, to provide a carbon source. The initial pH was
8.5, the average pH of the spring waters, and 1 mL L)1 of
vitamins (Wolfe’s mineral medium, Dworkin et al., 2006)
and 1 mL L)1 of trace elements (containing 0.52 g EDTA,
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D. F . GLEE SON e t al.
0.15 g FeCl2·4H2O, 7 mg ZnCl, 10 mg MnCl·4H2O,
6.3 mg H3BO3, 19 mg CoCl2·6H2O, 1.7 mg CuCl2·2H2O,
24 mg NiCl2·6H2O and 36 mg Na2MoO4·2H2O per 100 mL
deionized H2O) were also added to the solution.
Samples of mineral precipitates collected on the glacier were
used to inoculate the tubes, which were stored in the dark at
4 "C. Gradient tubes that contained visible yellow sulfur in
the overlayer after a period of about a month of growth were
then transferred to new gradient tubes to evaluate if S0 production continued. In subsequent transfers the plugs were
amended with 8 mM Na2S, rather than FeS, to remove Fe (II)
as another potential energy source for chemolithoautotrophic
growth. Autoclaved sediment from the spring deposits, collected in the field, was also added at 1% wt ⁄ vol to the plugs, to
ensure that potentially growth-limiting trace elements were
present in the cultures.
Salinity and sulfide concentrations were also increased from
initial salinities of 27–54& and 270&, while sulfide concentrations were increased from 8 to 16 mM and 80 mM in gradient tubes to evaluate the optimal conditions for S0
production as determined by visual inspection of tubes and
confirmed by microscopy. In addition, a subset of gradient
tubes was amended with pyruvate and formate (100 lM final
concentration) to test for sulfide oxidation during heterotrophic growth. Evaluation of whether sulfur production
increased with the addition of organic carbon was also determined by visual inspection of tubes and confirmed by microscopy.
Simplified cultures of sulfide-oxidizing bacteria
After 4 successive rounds of transfers in which the S0 was
microbially produced, the cultures were serially diluted in the
range of 10)1 to 10)7 in order to isolate the most abundant
members of the S0-generating consortia. Dilution series were
characterized by light and florescence microscopy after
1–3 months of growth. Successive rounds of serial dilutions
of S0 containing gradient tubes was conducted four times, and
DNA was extracted at various time-points (see below) to
determine the phylogeny of the consortia. In addition, some
of the S0 material was plated directly onto heterotrophic
media containing artificial seawater with yeast and peptone
and incubated in the dark at 4 "C. Plating onto autotrophic
sulfide media did not result in colony growth. Colonies of
varying morphology and pigmentation were then purified
through successive transfers on organic-rich ‘‘K’’-plates made
by adding 0.5 g yeast extract, 2 g peptone and 15 g granulated agar to 980 mL artificial seawater (containing 30 g
NaCl, 24 g MgSO4Æ7H2O, 3 g CaCl2Æ2H2O, 2 g KCl per
liter ultrapure H2O). After autoclaving, pre-sterilized solutions of 20 mL 1 M HEPES (pH 7.8) were added for a 20 mM
final concentration and 100 lL 1 M MnCl2 for a 100 lM final
concentration. Isolated colonies were grown in liquid
K-media, and DNA was extracted using the DNAEasy Kit
(Qiagen, Valencia, CA, USA) for 16S rRNA sequencing of
the isolates (see below).
All isolates obtained from heterotrophic plates were then
reintroduced to sulfide gradient tubes to evaluate whether
sulfide oxidation to S0 could be carried out in pure culture. Isolates were also introduced to agar-only tubes to determine if
isolates could be metabolizing the agar rather than the sulfide.
DNA extraction, cloning, and sequencing: enrichments, cocultures and isolates
Once mixed-culture S0-generating experiments had been
through five to six rounds of transfers, and microscopic
inspection had confirmed continuing sulfur production, cultures were considered to contain a semi-stable consortium.
DNA was then extracted from the highest-dilution gradient
tubes still producing S0. Sulfur-rich material from the gradient
tubes was first digested from the agar using the Qiaquick Gel
Extraction Kit from Qiagen. DNA was subsequently extracted
from the remaining material using the UltraClean extraction
kit (MoBio Inc., Carlsbad, CA, USA).
PCR reactions were carried out on the extracted DNA
using PCR primers 515F and 1074R (5¢-CACGAGCTGACGACAGCCAT-3¢) to amplify small-subunit rRNA gene
fragments for Bacteria. Due to the lack of Archaea in the environmental samples, no archaeal primers were used. Successful
PCR products with bands corresponding to the correct length
were cleaned with a Qiaquick PCR Purification Kit (Qiagen)
and cloned as above.
To sequence the clones, colony PCR reactions were conducted using M13F (5¢-GTAAAACGACGGCCAG-3¢) and
M13R (5¢-CAGGAAACAGCTATGAC-3¢) primers. Successful reactions were cleaned and the DNA concentration in each
sample was quantified from absorbance at 260 nm using a
Beckman-Coulter UV ⁄ VIS spectrophotometer. Samples were
then sent to Seqwright (Houston, TX, USA) to be sequenced
using M13F primers. The resulting sequences were processed
as above.
DNA was extracted from the purified isolates grown on
liquid K-media using the UltraClean extraction kit (MoBio
Inc.), and the DNA was amplified using 27F and 1492R primers. Full-length (!1500 bp) sequences were assembled using
a complement of 3–6 forward and reverse primers submitted
to Seqwright and processed as above.
Scanning electron microscopy (SEM) and energy dispersive
X-ray spectroscopy (EDS)
Scanning electron microscopy of environmental samples and
minerals produced in the culturing experiments was conducted using a JSM-6480LV (low vacuum) and JSM-7401F
(field emission) SEM at the Nanoscale Fabrication Laboratory
at the University of Colorado at Boulder. Sulfur-rich material
from the gradient tube experiments was fixed with glutaralde-
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Low temperature S0 biomineralization
hyde and dehydrated by exchanging solutions containing 50,
70, 75, 90, 95 and 100% ethanol for 30 min at each concentration. The samples were then applied to carbon filter paper,
air dried, and coated with 7 nm gold-palladium to reduce
charging of the samples during imaging. An Energy Dispersive X-ray Spectrometer was used to generate elemental analyses of the samples.
Genbank submission
16S rRNA gene sequences were submitted to Genbank under
accession numbers HM141098-HM141534. Sequences were
named according to sample name (BF64A and BF65B),
primer pair used (U – 515F, 1391R; B – 8F, 1492R) or cultivar (C – consortia, I – isolate), and sequence number.
RESULTS AND DISCUSSION
Geochemistry and mineralogy of the springs and deposits
The sulfide springs and associated sulfur deposits are dynamic
in nature, changing location and distribution from 1 year to
another. Field observations in 2000 and 2006 revealed extensive areas of yellow staining across the surface of the glacier.
On July 6, 2006 a large icing south of the toe of the glacier,
potentially built up by spring discharge, also had yellow staining with an areal extent of !0.12 km2. Satellite imagery of the
field site collected by the Hyperion instrument onboard Earth
Observing Satellite-1 allowed detection of the deposits in
2006, and in 2007 was utilized to track the disappearance of
these deposits over the course of the melt season (late Juneearly August). An onboard classification algorithm continued
to detect the presence of sulfur until snow obscured the site
again in late August 2007 (Gleeson et al., 2010). Field observations in other years observed only limited deposits in the
form of conical structures of sulfur, calcite and gypsum
(Grasby et al., 2003).
The measured spring water chemistry was saline
(7400 mg ⁄ L) and dominated by Na+, SO42) and Cl)
(Table 1). H2S at the spring outlet was measured at
142 mg L)1 (4.2 mM), the highest values yet reported in any
sulfur spring in Canada. The mineralogy of the supraglacial
deposits is dominated by S0, responsible for the yellow color
of the material of the ice, and calcite (Table 2). Although
quartz is also present in some of the samples, it is potentially
introduced to the system as wind-blown silt.
The accumulation of extensive S0 deposits is not predicted
to be thermodynamically stable under the highly oxygenated
and high pH conditions of 8 to 9 measured at Borup Fiord
(Gleeson et al., 2010). Sulfide (142 mg L)1) and sulfate
(1786 mg L)1) levels measured in spring waters provide a
lower limit on total sulfur in the system as the activities of
other potential intermediate sulfur species such as sulfite
(SO32)), thiosulfate (S2O32)) and polysulfides (Sn2)) were
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365
not measured. At the pH of 8–9 measured in the spring
waters, the dominant ion of sulfur is predicted to be either the
HS) ion or SO42), depending upon the Eh. High dissolved
Ca2+concentrations of 696 mg L)1 (58 mM) favor the precipitation of SO42) as gypsum at the ambient O2 levels if waters
are saturated, but we can only detect a limited amount of gypsum in the older dry deposits measured in previous years (Grasby et al., 2003), which may result from increased saturation
as the deposits dry.
Environmental community analysis
The microbial community composition of sulfur deposits
associated with the supraglacial spring at Borup Fiord was
analyzed with 16S rRNA gene sequencing. Clone libraries
were dominated by sequences representative of the Proteobacteria. Only one archaeal sequence and no eukaryal
sequences were identified after using PCR primers targeting
all three domains of life. Previous microbial community
analysis at the site identified members of the Proteobacteria, including microbes grouping within the Bulkholderiales
and Pseudomonadales and one sequence classified as a
Marinobacter sp. (Grasby et al., 2003). Sequences grouping
within the Burkholderiales and Pseudomonadales are also
identified in the clone libraries produced from environmental samples in the current study.
The 348 bacterial 16S rRNA sequences generated from
both environmental samples (Figs. 3–5) were dominated by
representatives of the b-Proteobacteria (!82% of total
sequences), of which !97% were most closely related to Ralstonia sp. Sequences grouping with Ralstonia sp. have been
identified in clone libraries produced from basal ice samples
collected from John Evans Glacier located on Ellesmere Island
(Skidmore et al., 2005). Ralstonia is a metabolically diverse
genus of the Burkholderia order, often prevalent in oxic,
Fig. 3 Bacterial diversity in clone libraries constructed from environmental
samples collected in 2006. Wedges represent phylum level distinctions, except
the Proteobacteria, which are divided into classes. ‘‘Other’’ refers to pooled
divisions representing <1% of each respective library. b-Proteobacteria in both
samples are dominated by Ralstonia.
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Fig. 4 Phylogenetic tree of the division Proteobacteria detected in 2006 environmental data, and in cultures inoculated from 2006 samples. Sequences obtained in
this study are in bold. Numbers following accession numbers indicate how many sequences grouped with each phylotype. Closely related organisms are found in many
sulfur-rich and cold environments.
metal-contaminated environments (Goris et al., 2001). Ralstonia are known to reduce iron in association with the oxidation of organic matter (Lin et al., 2007), can oxidize Fe(II)
under microaerophilic conditions (Swanner et al., 2011), and
can couple nitrate reduction to H2 oxidation (Zumft, 1997).
Ralstonia eutropha is a strictly respiratory lithoautotroph
whose genome has recently been shown to contain sulfur oxi-
dation (sox) genes (Cramm, 2009) although growth on thiosulfate was unsuccessful.
Other b-Proteobacteria present in clone libraries include
Comamonadaceae and Rhodocyclales. Microbes classified as
Comamonadaceae had been previously identified in samples
collected from the spring site by sequencing DGGE bands
(Grasby et al., 2003). Rhodocyclales include Rhodocyclus,
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Low temperature S0 biomineralization
367
Fig. 5 Phylogenetic tree of Bacteria excluding the division Proteobacteria within environmental samples and cultures. Sequences obtained in this study are in bold.
Numbers following accession numbers indicate how many sequences grouped with each phylotype. Closely related psychrophilic organisms are common, as are those
from saline environments.
which preferably grow photoheterotrophically under anoxic
conditions in the light with different organic substrates as carbon and electron sources. Chemotrophic growth is also possible under microoxic to oxic conditions in the dark, and
reduced sulfur compounds are not used as photosynthetic
electron donors by Rhodocyclus sp. (Imhoff, 2005).
c-proteobacteria representatives (!1% of total sequences)
included sequences related to Thiomicrospira sp. and Shewanella sp. One sequence was closely related (99% sequence
identity) to chemolithoautotrophic sulfur-oxidizing bacteria
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Thiomicrospira psychrophila and Thiomicrospira sp. which have
recently been shown to dominate clone libraries constructed
from sulfur streamers within cold sulfur springs on the neighboring island of Axel Heiberg Island (Neiderberger et al.,
2009). The low abundance of Thiomicrospira among the environmental sequences detected in 2006 may be linked to sulfide levels at Borup Fiord Pass, which are three orders of
magnitude higher than those of the springs on Axel Heiberg
Island. This may significantly impact the microbial community structure. The Shewanella representative was closely
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related to Shewanella sp. identified from cold, icy environments including the Blood Falls site in Antarctica (Mikucki &
Priscu, 2007).
e-proteobacteria representatives (3% of total sequences)
include sulfur-oxidizing bacteria that grouped with Sulfuricurvales and Sulfurovumales. Sulfuricurvales representatives
included sequences closely related (97% sequence identity) to
Sulfurimonas denitrifcans DSM 1251 (accession number
CP000153). Filamentous e-Proteobacteria including Sulfuricurvales and Sulfurovumales, related to those observed in
environmental data, are known to dominate sulfide-oxidizing
biofilms in warm sulfide-rich cave waters (Engel et al., 2003;
Macalady et al., 2008) and the e-Protobacterium Arcobacter
has been shown to oxidize sulfide, producing extracellular filaments of sulfur (Wirsen et al., 2002; Sievert et al., 2007).
Sequences not grouping within the Proteobacteria include
representatives of the Bacteroidetes, Chloroflexi, Actinobacteria and Firmicutes. Bacteroidetes representatives included
sequences closely related to Flavobacterium sp. identified at
Antarctic sites, including the Blood Falls site at Taylor Glacier
(Mikucki & Priscu, 2007). Sequences grouping with the Chloroflexi show close phylogenetic relationship to the Chloroflexi
sp. identified in the Guerrero Negro hypersaline mats (Spear
et al., 2003; Ley et al., 2006). Representatives of the Actinobacteria and Firmicutes also group with sequences identified
at cold, saline environments.
Bacterial diversity associated with the sulfur-rich springs at
Borup Fiord Pass as sampled in 2006 is broadly similar to
other cold, saline environments, including those of the other
High Arctic springs on Axel Heiberg Island (Perreault et al.,
2007, 2008) and subglacial flow from the Taylor Glacier in
the Antarctic (Mikucki & Priscu, 2007). Temperature and
chemistry of the spring waters likely act as the key drivers that
select for microbial community ecotype. Key differences from
A
B
the previous studies include the notable dominance of Ralstonia, for which a role in oxidative sulfur-cycling is currently
unknown.
Laboratory formation and biomineralization of S0
Initial sulfide gradient tubes inoculated with Borup Fiord sulfur deposits generated macroscopically visible cloudy white
clumps of S0 after a period of about 1 month. Mineralized
zones tended to grow outward from the line of inoculation
(Fig. 6) rather than in a band at a defined depth in the gradient tube. Production of S0 continued through successive
transfers, although the total amount of S0 was not as abundant
as found in the initial inoculations.
The sulfur-rich zones of the gradient tubes were examined
using differential interference contrast and fluorescence
microscopy. We typically observe abundant cells with a rod
morphology and semi-transparent, curved cylindrical filaments, some of which appear hollow while others are associated with spherical globules of sulfur along their lengths,
similar in appearance to the hydrated spherical colloid
described by Steudel (1989). Filaments vary from a few
microns to tens of microns in length (Fig. 7). Larger, rigid
sheaths are also observed, commonly mineralized with sulfur.
Extensively mineralized sheaths and filaments often exhibit
dense clusters of rounded and angular sulfur grains, which
commonly result in complete encrustation of the underlying
structure. The most typical morphologies include a central
mass of sulfur comprising one or more spheroidal mineral
aggregates, !5–50 lm in diameter, surrounded by narrow
filaments and biomineralized sheaths attached at one end of
their length and radiating out from the center of the structure
(Fig. 7). The morphologies of the spheroidal mineral aggregates have also been observed in studies investigating Borup
C
Fig. 6 (A) Gradient tube inoculated with Borup Fiord material shows the production of elemental sulfur in the form of cloudy, white, filamentous material. (B) False
colored DIC image shows filaments and mineralized sheaths in sulfur blooms from inoculated gradient tubes. (C) Abiotic precipitates in negative controls do not show
the same morphologies.
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Low temperature S0 biomineralization
369
A
Fig. 7 Further examples of sulfur structures
observed in enrichments. (A) DIC image shows that
cells are observed in close association with filaments. (B) Optical image shows full extent of a
sulfur structure. (C) Fluorescing cells against optical
background. (D) Sulfur crystals nucleating along
sheaths and filaments. (E) Sulfur globules deposited
along filaments.
B
C
D
E
Fiord field samples that confirmed these materials as sulfur
(Gleeson et al., in revision). Scanning electron microscopy
images demonstrate that the sheaths measure !1 lm across,
while the filaments are just a few hundred nm across (Fig. 8).
EDS spectra of the gold-coated samples confirm the deposited
material as sulfur while the sheaths themselves are dominated
by carbon.
Compositionally different but morphologically similar
structures have been reported by Benison et al. (2008), who
described clumps of organic bodies and sulfate crystals found
in evaporite minerals from Permian and modern acid saline
lakes as microbial remains. The similarities between our sulfur
structures and the ‘‘hairy blobs’’, despite compositional differ-
! 2011 Blackwell Publishing Ltd
ences, provide insight on the formation of S-mineralized
microbial structures, and strengthen the validity and application of these structures as combined morphological, mineralogical and organic biosignatures.
Optimal conditions for biological S0 production
During each transfer of the sulfide-oxidizing enrichments, the
consortia were subjected to serial dilutions up to 10)7. In each
experiment, HS) was provided as the sole electron donor, O2
as the sole electron acceptor (i.e., no nitrate was present), and
HCO3) as the sole carbon source. While agar was included in
early cultures to reduduce the rates of oxygen diffusion, and
370
D. F . GLEE SON e t al.
A
D
B
C
Fig. 8 (A–C) SEM images of extracellular sheaths and filaments at scales varying from 10 lm to 100 nm, (D) EDS spectra of material from image (B). These confirm
deposited grains as sulfur while indicating that the sheaths themselves comprise mainly carbon. NaCl crystals are also present in (A), an artifact of the preparation.
could have provided a carbon source for agar-degrading
micro-organisms, it was phased out during the course of the
experiments. In addition, agar-only tubes did not demonstrate growth, suggesting that our experiments did not select
for agar-dependent bacteria. In contrast, growth was observed
in all dilutions of Na2S gradient tubes, and S0 production was
variably maximized in the 10)3 to 10)5 dilutions, but rarely in
the most dilute. Our assumption is that full oxidation of sulfide to sulfate occurs to a greater degree when S0 is not
detected even though cell growth occurs, but this hypothesis
was not tested in this study. Instead, our focus was to deter-
mine the optimal conditions for microbial S0 production, and
to determine the phylogeny of the organisms in the stable
consortia.
Several variations in growth conditions were tested. First,
gradient tubes were amended with pyruvate and lactate
(100 lM final concentration), but we did not observe
enhanced S0 formation and these experiments were discontinued. Site spring waters with an average salinity of 7.4 g L)1
resulted in lower growth and sulfur production than our artificial EM seawater medium (27.5 g L)1 NaCl). Increased additions of NaCl to 2· and 10· EM also resulted in diminished
! 2011 Blackwell Publishing Ltd
Low temperature S0 biomineralization
371
growth and sulfur production, indicating that the consortia
are not highly halophilic. In addition, the sulfide concentrations in the plugs were increased from 4 mM to 8, 16 and
80 mM. Decreasing amounts of sulfur production and cell
growth occurred at increasing sulfide concentrations, and no
growth occurred at 80 mM NaS.
Increasing concentrations of NaCl and sulfide increased the
production of abiotic mineral precipitation within cultures, in
crystalline morphologies distinct from the sulfur structures
observed in our cultures. Abiotic precipitates at high sulfide
concentrations were much larger in size (!100 lm across)
than the structures observed in our cultures, with highly angular morphologies (Fig. 6). Previous studies of biogenic sulfur
have linked the monoclinic c-sulfur phase rosickyite with the
activities of microbial sulfide oxidation (Douglas & Yang,
2002; Douglas, 2004) and XRD measurements and SEM
images of Borup Fiord field samples have also revealed evidence of rosickyite (Gleeson et al., in revision). However, our
cultures did not yield enough sulfur to conduct similar XRD
analyses in this study.
S0 producing consortia
16S rRNA gene sequences of !500 bp were obtained from
extracted, cloned and sequenced DNA from 100 and 10)1
dilutions that produced S0 after five to six transfers. The consortia were dominated by the c-proteobacterium Marinobacter (Fig. 9). Other members of the consortia were closely
related to Shewanella, Loktanella, Pseudomonas, Rubrobacter,
and Sphingomonas spp. Consortia sequences show close phylogenetic relationships with environmental sequences identified from the field site as well as sequences identified from
other cold, saline environments. This indicates that we have
successfully enriched for stable consortia of environmentally
relevant organisms.
The respective roles of each cultivar are currently unknown,
but at least one organism detected has previously been implicated in sulfur-cycling. Loktanella salsilacus is an example of a
heterotrophic bacterium within the Rhodobacter group isolated from Antarctic lake environments (Van Trappen et al.,
2004), and closely related members of Loktanella sp. play
important roles in the cycling of organic and inorganic forms
of sulfur in marine environments. These roles include the degradation of dimethylsulfoniopropionate (DMSP) and oxidation of sulfite and thiosulfate (Gonzalez et al., 1999, 2000),
sulfur intermediates that may be present within the complex
redox cycle operating within the Borup Fiord spring system.
DNA was also extracted from a seventh generation S0generating gradient tube to evaluate whether the consortia
had changed. Conditions had been altered by the removal of
agar from the gradient tube over-layers to facilitate analysis
of the sulfur. The only organisms detected in the consortia
were Marinobacter sp. BF64A_C1 and Flavobacteria sp.
BF64A_C33 (also included in Fig. 9). While Flavobacterium
! 2011 Blackwell Publishing Ltd
Fig. 9 16S rRNA gene sequences from consortia within enrichments, dominated by Marinobacter.
was not previously represented in the 16S rRNA gene
sequences assembled for the consortia (likely due to too few
sequences), closely related Flavobacteria sequences are present
in the environmental data. It is possible that the change in
growth conditions to a fully liquid medium with higher oxygen concentrations may have been advantageous for this
organism, given its abundance in many freshwater and marine
environments (Kirchman, 2002).
Several organisms in the stable S0-generating consortia
were successfully isolated by plating the S0 from the gradient
tubes onto organic-rich media (K-plates containing yeast and
peptone), followed by the purification and sequence of colonies that varied in size and morphology. Pure cultures of
Shewanella sp. BF65B_I7, Loktanella sp. BF65B_I2, and
Sphingomonas sp. BF64A-I5 were obtained. In addition, Devosia sp. BF65B_I1 was also isolated (K. Wright, unpubl. data).
Devosia sp. BF65B_I1 was not detected as a major member of
the stable consortia during cloning and sequencing; however,
closely related Devosia have been detected in environmental
data. Marinobacter has not been isolated as a pure culture
from heterotrophic plates to date. However, Marinobacter
grows in mixed colonies with Shewanella sp. on the plates, and
when reintroduced into gradient tubes together, as determined by cloning and sequencing.
Each isolate, as well as the Marinobacter sp. BF64A_I1 ⁄
Shewanella sp. BF65B_I7 co-culture, was then used to inocu-
372
D. F . GLEE SON e t al.
late a set of sulfide gradient tubes to evaluate their capacity to
grow via sulfide oxidation in isolation. No sulfur production
or cell growth was observed in gradient tubes inoculated with
any of the isolates, including Shewanella sp. BF6 BF65B_I7.
However, the Marinobacter ⁄ Shewanella co-culture notably
did produce S0, cells, and biomineralized structures (supplemental data). DNA was extracted from the S0-mineralized
zones and limited cloning and sequencing again only detected
Marinobacter and Shewanella spp. Isolates were also used to
inoculate gradient tubes containing agar, but no sulfide, to
determine whether or not agar hydrolysis could be important
for growth; no cell growth was detected.
It is notable that the complex sulfur-generating consortia
present in the fifth and sixth round dilution series were found
to be dominated by Marinobacter, and that subsequently,
only experiments where Marinobacter was present continued
to generate sulfur and show both cell growth and sulfur biomineralization along sheaths and filaments. We suggest that
Marinobacter may be the key sulfide oxidizer in the system
(see below). Marinobacter sp. are common marine c-Proteobacteria, widely distributed throughout the water column, in
the deep ocean, and in Arctic and Antarctic ice (Brinkmeyer
et al., 2003; Shivaji et al., 2005; Zhang et al., 2008). Marinobacter sp. are well known as facultative heterotrophs with
motile rod shaped cells, and can use a wide variety of carbon
sources, including hydrocarbons, as their sole source of carbon and energy (Gauthier et al., 1992). The iron oxidation
capabilities of Marinobacter aquaeolei have recently been
described (Huu et al., 1999) and closely related strains have
been characterized as obligate chemolithoautotrophs
(Edwards et al., 2003). Marinobacter is typically halotolerant
or halophilic, while alkaliphic (M. alkaliphilus) (Takai et al.,
2005), psychrotolerant (M. maritimus) (Shivaji et al., 2005)
and psychrophilic (M. psychrophilus) (Zhang et al., 2008)
strains also exist. Marinobacter has not been shown to oxidize
sulfur species although it has been observed to grow in association with sulfate reducing bacteria (Sigalevich & Cohen,
2000). Recent results by Perreault et al. (2008) have shown
Marinobacter sp. NP40 to possess soxB, a gene involved in
thiosulfate oxidation in many Proteobacteria. Previous preliminary sequencing carried out for Borup Fiord samples by
Grasby et al. (2003) detected the presence of Marinobacter,
and the presence of this organism has been confirmed in environmental samples from 2006 (data not presented here).
From our work, it is possible that Marinobacter likely plays a
role in the oxidation of reduced S species.
While we do not have quantitative data on the abundance
of each species within our cultures, the dominance of Marinobacter in both early clone libraries and within simplified
enrichments and co-cultures points to this organism as the
likely sulfide oxidizer within our cultures. The close association of Marinobacter and Shewanella both on plates and in
gradient tubes may indicate that Shewanella plays a role
in cycling sulfur generated by Marinobacter, such as the
reduction of S0 described in Shewanella putrefaciens by Moser
& Nealson (1996). Shewanella and Flavobacteria could both
utilize organic materials generated by Marinobacter as their
heterotropic carbon source in their respective cultures, and
Flavobacteria are particularly well adapted to degrade biopolymers such as those in filament and sheath materials (Kirchman, 2002).
CONCLUSIONS
Studies of sulfide-oxidizing capabilities in the Proteobacteria
have traditionally focused on environments dominated by
mesophilic or thermophilic communities, such as caves and
hot springs (Spear et al., 2005, 2007). Pychrophilic sulfide
oxidation has been observed more recently in e-Proteobacteria (Skidmore et al., 2005), b-Proteobacteria (Sattley &
Madigan, 2006), and c-Proteobacteria (Knittel et al., 2005)
in a range of marine and subglacial environments. The
dominance of c-Proteobacteria related to Marinobacter, and
b-Proteobacteria related to Ralstonia, in our laboratory and
environmental data respectively, imply that these organisms
can thrive in S-dominated environments and their potential
for directly mediating sulfur oxidation reactions should be
closely evaluated.
Environmental clone libraries of the sulfur deposits are
dominated by Ralstonia sp., which are not currently known as
sulfur cyclers. These libraries also contain a range of organisms
known to play a role in the oxidation of sulfur compounds,
including Thiomicrospira, Sulfuricurvales and Sulfurovumales.
Moreover, our cultivation experiments indicate that members
of the microbial community present at Borup Fiord can grow
using sulfide as an energy source and catalyze the production
of S0. The organisms detected in our consortia, many of which
were also isolated, include Marinobacter, Shewanella, Loktanella, Sphingomonas, Pseudomonas, Flavobacterium, and
Devosia. These organisms were well represented in the environmental clone libraries described in this paper and previous
work by Grasby et al. (2003), and are considered relevant to
the Borup Fiord Pass spring system.
Marinobacter was found to dominate simplified sulfurgenerating cultures, and the generation of sulfur was not
observed in any experiments where Marinobacter was absent.
Furthermore, microscopic investigations (both in the dilutionto-extinction series and in the Marinobacter ⁄ Shewanella
cocultures) show that the sulfur is deposited in intimate association with microbial filaments and sheaths, indicating a
microbial control on morphology and distribution of the sulfur in our experiments. Abiotic controls for our experiments
did not show the same accumulations of sulfur; instead, limited abiotic precipitation only occurred in some high salinity
and sulfide experiments, and the precipitates were highly crystalline and distinct in size and morphology from those of the
inoculated experiments. Therefore, the distinct morphology of
the biogenic sulfur structures suggests that biomineralization
! 2011 Blackwell Publishing Ltd
Low temperature S0 biomineralization
associated with sulfide-oxidizing bacteria has the potential to
produce a morphological biosignature. S-mineralized microbial structures as morphological biosignatures could provide
targets for astrobiological investigations, linked to macroscale
mineral deposits that could be visualized and interpreted from
distance. The existence of mineral biosignatures of this nature
in a terrestrial setting informs the search for biosignatures in
other locations in the solar system, such as at Mars and at the
icy surface of Europa.
ACKNOWLEDGMENTS
The authors acknowledge Benoit Beauchamp at the University of Calgary for collaboration in the field and Katherine
Wright at the University of Colorado for collaboration in the
laboratory. Fieldwork was supported by the Canadian Polar
Continental Shelf Project, The Planetary Society and a Lewis
and Clark Field Scholarship. Laboratory work was conducted
with funding from the Director’s Discretionary Fund of
the NASA Astrobiology Institute and the David and Lucille
Packard Foundation. Portions of this work were carried out at
the Jet Propulsion Laboratory, California Institute of Technology, under a contract with the National Aeronautics and
Space Administration.
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SUPPORTING INFORMATION
Additional Supporting Information may be found in the
online version of this article:
Figure S1. Follow-up imaging on sulfur structures within enrichments and
co-cultures of Marinobacter and Flavobacteria.
Figure S2. Follow-up imaging of sulfur structures within co-cultures of Marinobacter and Shewanella, showing central masses comprising one or more
spherical mineral aggregates, curved filaments and wholly or partially mineralized rigid sheaths.
Figure S3. Different stages in formation of sulfur structures by consortia.
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