1
Supplementary materials
2
3
METHODS
4
5
Total Nucleic Acids Extraction
6
The same total nucleic extraction protocol (Dempster et al., 1999) was followed for all of
7
the samples. The quarter of each filter dedicated to molecular work was stored in separate
8
microcentrifuge tubes at -80°C in a cetyl trimethylammonium bromide (CTAB) solution
9
consisting of 100 mM TrisHCl at pH 8.0, 1.4 M NaCl, 2% (w/v) CTAB, 1.0% polyvinyl
10
pyrrolidone, and 20 mM Ethylenediaminetetraacetic acid. The filters and solution were
11
thawed at room temperature, and 0.4% (v/v) beta-mercaptoethanol was added to each
12
tube. The samples were vortexed briefly and incubated at 65°C for 15 min with
13
occasional inversion. The samples were cooled to room temperature, and an equal
14
volume of chloroform/isoamyl alcohol (24:1) was added, briefly mixed, and incubated at
15
room temperature for 20 min. The aqueous and the organic phases were separated by
16
centrifugation at 12,500 rpm for 20 min at 4°C. The aqueous layer was transferred to a
17
new tube, an equal volume of chloroform/isoamyl alcohol was added and briefly mixed,
18
and the phases were separated by centrifugation for 5 min. The aqueous layer was again
19
moved to a new tube, and ½ volume 5 M NaCl and 1 volume isopropanol were added and
20
briefly mixed. The samples were frozen at -80°C for at least 2 hours and then thawed and
21
centrifuged at 4°C for 45 min. The pellet was washed with 70% ethanol and resuspended
22
in 50 µL of RNase free water.
23
24
PCR, RT-PCR and Cleaning
25
All of the PCR reactions for the generation of clone libraries used the same bacterial 16S
26
rRNA primers-- 8F (5’-AGRGTTTGATCCTGGCT CAG-3’) and 1492R (5’-
27
CGGCTACCTTGTTACGACTT-3’; Teske, et al., 2002). Each PCR reaction contained 1
28
µL of DNA template, 2.0 µL of each primer solution (10 µM each), 0.25 µL of
29
SpeedSTAR Polymerase (Takara, Shiga, Japan), 2.0 µL of deoxynucleotide triphosphate
30
(2.5 mM of each dATP, dCTP, dGTP and dTTP) and 2.5 µL of 10X Fast Buffer I; the
31
volume was adjusted to 25 µL with molecular biology-grade water. Conditions for the
1
32
PCR in the Bio-Rad iCycler (Hercules, CA, USA) were as follows: heat activation at
33
94°C for 2 min, followed by 25 cycles consisting of 10 s denaturation at 98°C, 15 s
34
primer annealing at 60°C, and 20 s elongation at 72°C. The PCR terminated with a 72°C
35
extension at 10 min.
36
37
For the production of the 16S rRNA clone libraries, an aliquot of the extracted total
38
nucleic acids was treated with RNase-free DNase I using the Turbo DNA-free kit
39
(Ambion, Austin, TX, USA). The DNA-free samples (1 µL) were used in a 25 µL reverse
40
transcriptase chain reaction (RT-PCR) with a Real-Time One-Step RNA PCR Kit,
41
Version 2.0 (Takara) with 12.5 µL of 2X One Step RNA PCR Buffer, 2.0 µL of each
42
primer solution (of 10 µM), 0.5 µL of RNase inhibitor, 0.5 µL of TaKaRa Ex Taq HS,
43
and 0.5 µL of Reverse Transcriptase XL (AMV). Conditions for the RT-PCR in the Bio-
44
Rad iCycler were as follows: reverse transcription at 42°C for 15 min, reverse
45
transcriptase inactivation at 95°C for 2 min, followed by 25 cycles consisting of 20 s
46
denaturation at 98°C, 25 s primer annealing at 60°C, and 1 min elongation at 72°C. The
47
RT-PCR terminated with an extension at 72°C for 10 min. The coastal PCR (DNA) and
48
RT-PCR (cDNA) products were cleaned using a Wizard SV Gel and PCR Clean-Up
49
system (Promega, Madison, WI, USA) using either the gel purification or the PCR
50
product purification protocols provided with the kit. The offshore DNA and cDNA were
51
cleaned using an UltraClean GelSpin DNA Extraction Kit (MoBio Laboratories, Solana
52
Beach, CA, USA). Samples that gave >30% chimeric results were re-processed using a
53
reconditioning PCR (Thompson et al., 2002). The PCR product (1 µL) was used in a new
54
PCR reaction with new reagents for 3 more cycles, and the resulting PCR product was
55
not cleaned before cloning.
56
57
The clean PCR and RT-PCR products were cloned into chemically competent E. coli
58
cells using a TOPO TA Cloning Kit that contained TOP 10 cells and either a pCR 2.1-
59
TOPO vector or a pCR 4.0 TOPO vector (Invitrogen, San Diego, CA, USA). Sanger
60
sequencing of the bacterial colonies was performed in the forward and reverse direction
61
by GENEWIZ, Inc. (South Plainfield, NJ) on an ABI 3730xl sequencer using universal
62
primers M13F(-21) and M13F(-47). Contiguous sequences were constructed and edited
2
63
using Sequencher (Genecodes, Ann Arbor, MI, USA). Clean, full-length sequences were
64
sent to greengenes.lbl.gov for chimera checking using the default settings of Bellerophon
65
version 3 (Huber et al., 2004).
66
67
RESULTS AND DISCUSSION
68
69
Surface/ Subsurface, DNA/RNA, and Particle-Associated Partitioning
70
The OTUs that represented five or more sequences (28 OTUs total) are shown in the heat
71
map (Fig. S3) with the percentage of each library for which they account. For all OTUs
72
that were represented in more than one library (57 OTUs total), 95% showed partitioning
73
into surface waters (both coastal and offshore) and deeper (mid-depth and bottom)
74
waters. Of the top 28 OTUs, 18% were exclusively found in particle-associated or free-
75
living libraries, and 43% were exclusively found in DNA or RNA libraries. Five OTUs
76
were only found in DNA libraries: the ‘Candidatus Pelagibacter ubique’ (Fig. S5a),
77
Actinobacteria gp. OCS155 (Fig. S5d), SAR11-Surface 1 (Fig. S5a), Micromonas (Fig.
78
S5d), and SAR86 (II) (Fig. S5b) groups. Seven OTUs were only found in RNA libraries:
79
Thalassospira (Fig. S5a), the Eastern North Pacific Ocean SUP05 cluster (Fig. S5b), the
80
MGB / SAR324 Clade – Group II (Fig. S5d), Sulfitobacter (Fig. S5a), Arctic BD96
81
Group (Fig. S5b), Flavobacterial Marine Group NS9 (Fig. S5c), and the SAR324-
82
Chesapeake Bay group (Fig. S5d).
83
84
Of the top 28 most common OTUs, only two were found in both a surface (coastal or
85
offshore) library and a mid-depth or bottom depth offshore library. One was an OTU
86
closely related to Pseudoalteromonas undia (Fig. S5b), of which most strains are capable
87
of protease, lipase, and amylase activity, and possibly galactosidase activity (Yu et al.,
88
2009). The other OTU in this category grouped with Hyphomonas (Fig. S5a), some
89
members of which have been characterized by an approximate 0.9 µm diameter and a
90
reproduction method of budding from the tip of a hypha that is up to three times longer
91
than the parent cell (Weiner et al., 1985). The OTU was found exclusively in the particle-
92
associated offshore surface and mid-depth libraries.
93
3
94
Coastal and Offshore Surface Partitioning
95
Several OTUs were preferentially found in the coastal libraries. Two of these OTUs are
96
closely related to a Roseobacter clone (Fig. S5a) and to a Marine Group B / SAR 324
97
Clade clone (Fig. S5d) recently identified in the Chesapeake Bay (Kan et al., 2008). An
98
OTU closely related to the “Candidatus Pelagibacter ubique” strain HTCC1002 (Fig.
99
S5A), a coastal SAR11 isolate that carries out glycolysis (Schwalbach et. al, 2010), was
100
found in the coastal and offshore station libraries. It accounted for a greater percentage of
101
the coastal libraries than the offshore libraries. An OTU belonging to the OCS155 clade
102
of Actinobacteria (Fig. S5e) represented more of the coastal than the offshore surface
103
libraries. This deeply branching group has been found in the Sargasso Sea, the
104
continental shelf waters off North Carolina, the Northeastern Pacific Ocean, and the
105
Columbia River mouth and estuary (Rappé et al., 2000). An OTU in the SAR116 group
106
(Fig. S5a) was exclusively found in the coastal station libraries. The genome of the first
107
cultured representative of this group, “Candidatus Puniceispirillum marinum” IMC1322,
108
was recently sequenced. The genome suggests that the group is composed of metabolic
109
generalists in ocean nutrient cycling (Oh et al., 2010). The Flavobacteria Marine Group
110
NS9 OTU (Fig. S5c) was preferentially found in the coastal libraries. A SAR11 Surface
111
1b group (Fig. S5a; Carlson et al., 2008) was found exclusively at the surface offshore, as
112
were the OTUs from the genera Sulfitobacter (Fig. S5a), Prochlorococcus (Fig. S5e), and
113
Thalassospira (Fig. S5a). One OTU was noticeably not preferentially found in the coastal
114
or offshore surface libraries. This OTU was most closely related to Synechococcus sp.
115
RS9905 (Fig. S5d), a member of clade III; this clade differs from the other clades
116
because its members are motile (Toledo et al., 1999). The Synechococcus OTU was
117
found in the coastal and offshore surface libraries.
118
119
Notable Bacterial Groups
120
The two most common OTUs were both Gammaproteobacteria that were only found at
121
the mid-depth and bottom depths offshore. The first of these (Fig. S5b) was represented
122
by 73 sequences (10.1% of the 723 total sequences from all libraries) and grouped with
123
the genus Marinomonas, but did not match cultured isolates. A closely-related,
124
uncultured species (AY028196) was identified in bacterioplankton samples exposed to
4
125
diatom detritus (Bidle & Azam, 2001). The second-most observed OTU (Fig. S5b)
126
grouped with Balneatrix. It was closely related to an uncultured endosymbiont of a cold-
127
seep Mytilidas sp. (Duperron et al., 2008). An Alphaproteobacteria OTU grouping with
128
the genus Thalassospira (Fig. S5a) was only found in a surface offshore library. It
129
represented 51% of the free-living offshore surface RNA library and a small portion (5%)
130
of the particle-associated surface RNA library. This OTU’s closest cultured
131
representative is Thalassospira tepidiphila, a motile, facultatively anaerobic polycyclic
132
aromatic hydrocarbon-degrading bacterium (Kodama et al., 2008).
133
134
A gammaproteobacterial OTU (Fig. S5b) within the genus Oceanospirillales was found
135
exclusively in the four offshore mid-depth libraries, and is very closely related to
136
uncultured representatives (HM587889) of the DWH Oceanospirillales found in Gulf of
137
Mexico Deepwater Horizon deep hydrocarbon plume (Hazen et al., 2010; Mason et al.
138
2012). Interestingly, the mid-depth water mass might contain hydrocarbons from natural
139
seeps, as was observed for the Subtropical Underwater in the southwest North Atlantic
140
(Harvey et al., 1979; Requejo & Boeh, 1985). Because the subtropical underwater of the
141
southwest North Atlantic extends into the Gulf of Mexico, it constitutes a potential source
142
for the Oceanospirillales-related bacteria that responded quickly to the availability of
143
dissolved hydrocarbons in the oil-polluted water column (Hazen et al. 2010). A second
144
Oceaniserpentilla cluster was distinct from the first Oceaniserpentilla OTU, but was
145
distributed among the libraries similarly.
146
147
One OTU grouped with Marine Group B/SAR324 Clade Group II (Fig. S5d; Brown and
148
Donachie, 2006). Group II has been found globally distributed in cold waters (<16°C;
149
Wright et al., 1997), and enzyme sequencing has suggested that it is important in
150
dissolved organic phosphorus cycling (Brown & Donachie 2006). It represented a
151
considerable portion of the free-living offshore bottom RNA library (20%), and is also
152
found in the free-living offshore mid-depth RNA library (3%). Only the Synechococcus
153
sp. RS9905 OTU (Fig. S5d) was found in the coastal and offshore surface RNA and
154
DNA libraries.
155
5
156
One of the most dominant marine bacterial groups, the SAR11 cluster, was abundant in
157
the DNA coastal and offshore libraries and noticeably missing from the corresponding
158
RNA libraries (Fig. 2 and S5a). There are no obvious methodological reasons for this
159
discrepancy. The total nucleic acid extraction protocol extracted both DNA and RNA at
160
the same time, and the same primers were used for both DNA and RNA libraries.
161
Additionally, Moeseneder et al. (2005) found the SAR11 group using T-RFLP of both
162
RNA and DNA, indicating that preferential SAR11 RNA degradation is unlikely. The
163
SAR11 group simply may not have been very active in our samples, despite evidence that
164
SAR11 bacteria can be relatively large and active off the North Carolina coast in April
165
(Malmstrom et al., 2004). One other study, to the knowledge of the authors, has
166
demonstrated low activity in SAR11 bacteria. In the Arctic, approximately 2% - 50% of
167
SAR11 bacteria showed uptake of glucose and amino acids using micro-FISH, and even
168
fewer were actively taking up ATP; other groups such as Roseobacter,
169
Alphaproteobacteria, Betaproteobacteria, and Gammaproteobacteria had higher
170
percentages of active bacteria (Alonso-Sáez et al., 2008). The difference in SAR11
171
representation between DNA and RNA surface water clone libraries does not cause the
172
enhanced DNA and RNA division in the surface compared to the mid-depth and bottom
173
libraries, as shown by FastUniFrac analysis excluding SAR11 (results not shown).
174
175
Linkage between community composition and enzymatic activity.
176
Comparing the potential of the microbial community to respond to addition of specific
177
substrates with microbial community composition is a challenge for the experimental
178
design of enzyme assays. Here we chose to assess the potential of living cells, gravity
179
filtered onto different filter sizes, in time course experiments. After filtration, and
180
incubation of the filter pieces in artificial seawater, initially only the activity of enzymes
181
attached to cells (or detritus) on the filter is measured. Once hydrolytic enzymes are
182
synthesized, they most likely remain active for several days or longer (Steen and Arnosti,
183
2011); therefore, hydrolysis rates will, over the time course of an incubation, partially
184
uncouple from the changing composition of the microbial community. Microbial
185
community change during incubation time will depend on the average generation time of
186
the community, and can modulate the spectrum of enzyme activities. The enzyme
6
187
activities that dominate at any time point represent a time-integrated microbial
188
community response over the length of the incubation. Conceptually, microbial
189
community composition and potential hydrolysis rates are linked most directly at the start
190
of an incubation, when the clone library composition reflects the “starter” bacterial
191
community that had synthesized the enzymes whose integrated activity spectrum is
192
assessed at the first time point after filtration (Fig. 1). In contrast, enzymes whose
193
activities are detected later in the incubation likely reflect the development of enzymatic
194
activity by enzyme production (Fig. 1, Fig. S2).
195
196
The Shannon Index was used as previously described (Steen et al., 2010) to evaluate the
197
evenness of the hydrolytic capabilities of a community, analyzed after 2 days incubation,
198
as well as using the maximum rates that were observed over the timecourse of incubation
199
(see Fig. S2). Using maximum hydrolysis rates, the evenness of hydrolysis rates for
200
particle-associated and unfiltered water decreased (according to significant differences)
201
in the following order: coastal, offshore mid-depth, surface, and bottom waters (Fig. S8).
202
For the free-living community with maximum hydrolysis rates, the Shannon index was
203
highest at the coastal station and did not vary with depth offshore. Similar results were
204
obtained for analysis using the hydrolysis rates measured on day 2 (Fig. S8). The
205
diversity/evenness of hydrolysis rates thus did not map directly to diversity of OTUs at
206
the activity levels and phylogenetic levels we examined.
207
The contrast between the community diversity and hydrolytic capability is particularly
208
notable for the offshore bottom water communities. Though more diverse than the mid-
209
depth communities (Fig. 2 and S4), the bottom communities hydrolyzed a narrower
210
spectrum of substrates (Fig. 1), and evenness of hydrolysis rates were also generally
211
lower, especially for the particle-associated community (Fig. S8). A similar disconnect is
212
apparent in the comparison of community diversity and hydrolytic capability in coastal
213
and offshore samples. The microbial community at the coastal station hydrolyzed all six
214
fluorescently-labeled substrates at higher rates than at the offshore surface station, where
215
fewer substrates were hydrolyzed (Fig. 1). These broad capabilities and higher rates of
216
hydrolysis at the coast are not matched by greater bacterial 16S rRNA clone library
217
diversity. Detailed understanding of the relationship between community composition
7
218
and hydrolytic capabilities will likely require genomic and proteomic studies to identify
219
the range of enzymes produced by specific phylogenetic groups (e.g., Wegner et al.
220
2013), since extracellular enzymatic capabilities of bacteria are likely differentiated at
221
fine-grained phylogenetic scales (Zimmerman et al. 2013).
222
223
Rationale for Time Course Experiments of Extracellular Enzymatic Activities
224
The clone libraries constructed using 16S rRNA genes were intended to assess the
225
bacterial community that potentially participates in polysaccharide degradation. These
226
bacteria may be more likely to hydrolyze a specific substrate if they have the necessary
227
enzymatic tools. If the active community does not possess these tools, then an inactive
228
but relatively abundant group that is capable of hydrolysis may be activated.
229
Alternatively, a relatively rare group with hydrolytic capability could increase in
230
abundance. Because of these different possibilities, maximum rates of hydrolysis most
231
likely do not correspond to the activity of initially active groups represented in the RNA
232
libraries. However, the time course of hydrolysis can give important clues about the
233
extent to which individual metabolic and/or community composition restructuring must
234
take place for hydrolysis to occur. For example, despite the great diversity across all
235
RNA libraries, laminarin was hydrolyzed quickly in all samples, supporting the
236
hypothesis that a wide range of marine bacteria can hydrolyze laminarin. Mixed
237
community incubations more accurately portray the hydrolytic capabilities of natural
238
communities than experiments with individual isolates. Given the time scale of
239
incubation, the response of a microbial community to a substrate could include enzyme
240
induction, cellular growth, and changes in community composition. However, it is
241
important to remember that even if a community hydrolyzes substrate quickly, the
242
fraction of the metabolically active community performing the hydrolysis is unknown.
243
244
Time Courses of Hydrolysis
245
Among substrates hydrolyzed by particle-associated and free-living microbial
246
communities of both the coastal station and the offshore station surface site, two
247
substrates had noticeably different patterns of hydrolysis with time: chondroitin and
248
xylan (Fig. S2). These different patterns likely reflect differing time-scales of response by
8
249
the microbial communities. A rapid response suggests either the widespread presence of a
250
gene(s) for the appropriate enzyme(s) among diverse members of the community, and/or
251
the presence of the gene(s) for the enzyme among abundant members of the community.
252
A slow response (increasing hydrolysis rates with time) likely indicates an increased
253
growth response by a rare group capable of hydrolyzing the substrate, induction of
254
enzymes among such community members, and/or a shift in metabolism as low
255
concentrations of natural substrates in the incubation are exhausted.
256
257
Laminarin hydrolysis rates were always maximal at the first time point, fitting the pattern
258
discussed above for rapid response. In contrast, chondroitin only reached its maximum
259
hydrolysis rate by the first time point once, in the particle-associated coastal station
260
sample. For all other samples, maximum chondroitin hydrolysis rates occurred after an
261
initial time lag, a response consistent with enzyme induction or slow growth. An
262
experiment involving pre-exposure of a microbial community to chondroitin in fact
263
suggested that enzyme induction can occur for chondroitin on the time scale of hours to
264
days (Arnosti, 2004). Pure culture experiments with several organisms have also shown
265
inducibility of chondroitin- hydrolyzing enzymes (Lipeski et al., 1986; Shain et al.,
266
1996).
267
268
The pattern of xylan hydrolysis over time differed between the coastal and offshore
269
locations more than that of any other substrate. Xylan hydrolysis was still increasing on
270
day 13 in the particle-associated and free-living coastal incubations, while it was steady
271
in the particle-associated and free-living offshore station surface samples (Fig. S2). The
272
coastal pattern is consistent with hydrolysis performed by an initially small portion of the
273
population that grows over the time course of the incubation. The time course of xylan
274
hydrolysis also differed with depth more than that of other substrates. Xylan hydrolysis
275
was delayed in the bottom offshore station samples compared with the hydrolysis by the
276
mid-depth and surface samples (Fig. S2). This delay is consistent with either induction by
277
the substrate or the growth of a hydrolyzing population that was initially rare.
278
9
279
The time course of hydrolysis varied between the particle-associated and free-living
280
fractions for some substrates, even though particle-associated and free-living
281
communities nearly always hydrolyzed the same substrates. The most sensitive
282
information regarding differences in bacterial composition and enzymatic activity may be
283
found in the time course of the hydrolysis of these substrates. Fucoidan hydrolysis at the
284
coastal station occurred quickly in the free-living sample, but began on day 8 and was
285
still increasing by day 13 in the particle-associated sample. The opposite was true for
286
arabinogalactan and chondroitin hydrolysis, both of which reached their maximum rates
287
of hydrolysis earlier in the particle-associated coastal station sample than in the free-
288
living sample. The offshore stations had fewer differences between particle-associated
289
and free-living hydrolysis rate patterns than the coastal station; the most notable
290
difference was the delayed chondroitin hydrolysis in the particle-associated compared to
291
the free-living fraction in the bottom site offshore station sample.
292
293
294
REFERENCES
295
296
Alonso‐Sáez L, Sánchez O, Gasol Josep M, Balagué V, Pedrós‐Alio C. (2008).
297
Winter‐to‐summer changes in the composition and single‐cell activity of near‐surface
298
Arctic prokaryotes. Environ Microbiol 10:2444-2454.
299
300
Arnosti C. (2004). Speed bumps and barricades in the carbon cycle: substrate structural
301
effects on carbon cycling. Mar Chem 92:263–273.
302
303
Bidle KD, Azam F. (2001). Bacterial control of silicon regeneration from diatom detritus:
304
significance of bacterial ectohydrolases and species identity. Limnol Oceanogr 46:1606–
305
1623.
306
307
Brown MV, Donachie SP. (2006). Evidence for tropical endemicity in the
308
Deltaproteobacteria Marine Group B/SAR324 bacterioplankton clade. Aquat Microb Ecol
309
46:107–115.
10
310
311
Carlson CA, Morris R, Parsons R, Treusch AH, Giovannoni S. J, Vergin K. (2008).
312
Seasonal dynamics of SAR11 populations in the euphotic and mesopelagic zones of the
313
northwestern Sargasso Sea. ISME J 3:283–295
314
315
Dempster EL, Pryor KV, Francis D, Young JE, Rogers HJ. (1999). Rapid DNA
316
extraction from ferns for PCR-based analyses. Biotechniques 27:66.
317
318
Duperron S, Halary S, Lorion J, Sibuet M, Gaill F. (2008). Unexpected co-occurrence of
319
six bacterial symbionts in the gills of the cold seep mussel Idas sp. (Bivalvia: Mytilidae).
320
Environ Microbiol 10:433–445.
321
322
Hazen TC, Dubinsky EA, DeSantis TZ, Andersen GL, Piceno YM, Singh N, et al,
323
(2010). Deep-sea oil plume enriches indigenous oil-degrading bacteria. Science 330:204-
324
208.
325
326
Harvey G, Requejo A, McGillivary P, Tokar J. (1979). Observation of a subsurface oil-
327
rich layer in the open ocean. Science 205:999–1001.
328
329
Huber T, Faulkner G, Hugenholtz P. (2004). Bellerophon: a program to detect chimeric
330
sequences in multiple sequence alignments. Bioinformatics 20: 2317-2319.
331
332
Kan J, Evans SE, Chen F, Suzuki MT. (2008). Novel estuarine bacterioplankton in rRNA
333
operon libraries from the Chesapeake Bay. Aquat Microbial Ecol 51: 55-66.
334
335
Kodama Y, Stiknowati LI, Ueki A, Ueki K, Watanabe K. (2008). Thalassospira
336
tepidiphila sp. nov., a polycyclic aromatic hydrocarbon-degrading bacterium isolated
337
from seawater. Int J Syst Evol Microbiol 58: 711-715.
338
11
339
Lipeski L, Guthrie EP, O’Brien M, Kotarski SF, Salyers AA. (1986). Comparison of
340
proteins involved in chondroitin sulfate utilization by three colonic Bacteroides species.
341
Appl Environ Microbiol 51:978-984.
342
343
Malmstrom RR, Kiene RP, Cottrell MT, Kirchman David L. (2004). Contribution of
344
SAR11 bacteria to dissolved dimethylsulfoniopropionate and amino acid uptake in the
345
North Atlantic Ocean. Appl Environ Microbiol 70:4129-4135.
346
347
Marshall KT, Morris RM. (2012) Isolation of an aerobic sulfur oxidizer from the
348
SUP05/Arctic96BD-19 clade. ISME J. 7:452-455.
349
350
Mason OU, et al. (2012) Metagenome, metatranscriptome and single-cell sequencing
351
reveal microbial response to Deepwater Horizon oil spill. ISME J. 6:1715-1727.
352
353
Moeseneder MM, Arrieta JM, Herndl GJ. (2005). A comparison of DNA-and RNA-based
354
clone libraries from the same marine bacterioplankton community. FEMS Microb Ecol
355
51:341–352.
356
357
Oh HM, Kwon KK, Kang I, Kang SG, Lee JH, Kim SJ, Cho JC. (2010). Complete
358
genome sequence of “Candidatus Puniceispirillum marinum” IMCC1322, a
359
representative of the SAR116 clade in the Alphaproteobacteria. J Bacteriol 192: 3240–
360
3241.
361
362
Rappé MS, Vergin K, Giovannoni SJ. (2000). Phylogenetic comparisons of a coastal
363
bacterioplankton community with its counterparts in open ocean and freshwater systems.
364
FEMS Microbiol Ecol 33:219–232.
365
366
Requejo A, Boehm P. (1985). Characterization of hydrocarbons in a subsurface oil-rich
367
layer in the Sargasso Sea. Mar Environ Res 17: 45–64.
368
369
Schlitzer R. (2002). Ocean Data View, http://www.awi-bremerhaven.de/GEO/ODV.
12
370
371
Schwalbach MS, Tripp HJ, Steindler L, Smith DP, Giovannoni SJ. (2010). The presence
372
of the glycolysis operon in SAR11 genomes is positively correlated with ocean
373
productivity. Environ Microbiol 12: 490–500.
374
375
Shain H, Homer KA, Beighton D. (1996). Degradation and utilisation of chondroitin
376
sulphate by Streptococcus intermedius. J Med Microbiol 44:372-380.
377
378
Steen AD, Arnosti C (2011) Long lifetimes of β-glucosidase, leucine aminopeptidase,
379
and phosphatase in Arctic seawater. Marine Chem 123: 127-132.
380
381
Steen AD, Ziervogel K, Arnosti C. (2010). Comparison of multivariate microbial datasets
382
with the Shannon index: An example using enzyme activity from diverse marine
383
environments. Org Geochem 41:1019-1021.
384
385
Teske, A, Hinrichs KU, Edgcomb V, de Vera Gomez A, Kysela D, Sylva SP, Sogin ML,
386
and Jannasch HW. (2002). Microbial Diversity of Hydrothermal Sediments in the
387
Guaymas Basin: Evidence for Anaerobic Methanotrophic Communities. Appl Environ
388
Microbiol 68: 1994-2007.
389
390
Thompson JR, Marcelino LA, Polz MF. (2002). Heteroduplexes in mixed-template
391
amplifications: formation, consequence and elimination by “reconditioning PCR. Nucleic
392
Acids Res 30:2083.
393
394
Toledo G, Palenik B, Brahamsha B. (1999). Swimming marine Synechococcus strains
395
with widely different photosynthetic pigment ratios form a monophyletic group. Appl
396
Environ Microbiol 65:5247–5251.
397
398
Weiner RM, Devine RA, Powell DM, Dagasan L, Moore RL. (1985). Hyphomonas
399
oceanitis sp.nov., Hyphomonas hirschiana sp. nov., and Hyphomonas jannaschiana sp.
400
nov. Int J Syst Bacteriol 35: 237–243.
13
401
402
Yu Y, Li H, Zeng Y, Chen B. (2009). Extracellular enzymes of cold-adapted bacteria
403
from Arctic sea ice, Canada Basin. Polar Biology 32: 1539–1547.
404
405
14
406
Figure legends
407
408
Figure S1. Sampling locations and water column profiles. a) Coastal and offshore
409
station locations graphed using Ocean Data View (Schlitzer, 2002). b). CTD profiles.
410
Line 1= temperature; Line 2 (bold) = salinity; Line 3 (grey) = oxygen concentration; and
411
Line 4 (thin) = beam transmission. Horizontal dotted lines indicate the surface, mid-
412
depth, and bottom sampling depths.
413
414
Figure S2. Time course of hydrolysis of six different substrates by coastal and offshore
415
communities. Error bars show deviation between two replicates. PA = Particle-
416
associated; FL = Free-living.
417
Figure S3. Distribution heat map of OTUs among all 16S rRNA and rRNA gene
418
clone libraries. First letter and number of clone names indicate the tree
419
(g=Gammaproteobacteria, a = Alphaproteobacteria, o = Other, b = Bacteroidetes) and
420
position in tree. For example, clone g16Ys0Os112 is found in the Gammaproteobacteria
421
tree. It is the 16th clone from the top, and it is found in the tree as g16Ys0Os112. ALL
422
numbers represent percentage of each library the OTU composes. First bar = 73 seqs. PA
423
= Particle-associated library; FL = Free-living library; D = DNA library; R = RNA
424
library. OTU definition = 0.03. (DWH Oceanospirillales indicates the clone that showed
425
99.6% nucleotide similarity to a prominent Oceanospirillales clone in the Deepwater
426
Horizon subsurface hydrocarbon plume (Hazen et al., 2010).
427
428
Figure S4. Rarefaction curves for all clone libraries. FL = Free-living; PA = Particle-
429
associated. OTU definition = 0.03.
430
431
Figure S5. Phylogenetic trees based on near-complete 16S rDNA and rRNA sequences,
432
and calculated by empirically optimized distances following the General Time Reversible
433
plus Invariant sites plus Gamma (GTR+I+G) distribution model. Bootstrap support was
434
obtained by 200 replicates using the same model. The trees show the relationships
435
between all clones in all sixteen 16S rRNA and rDNA clone libraries, separated into the
436
following groups: a) Alphaproteobacteria, b) Gammaproteobacteria, c) Bacteroidetes, d)
15
437
Delta-, Epsilon- and Betaproteobacteria, e) Other bacterial phyla. Asterisks* indicate
438
clones that represent multiple similar sequences; numbers of sequences are given in
439
parentheses.
440
441
Figure S6. Cell counts and hydrolysis rates. Hydrolysis rates in whole water (same
442
data as in Fig. 1a), plotted along with cell counts in whole water.
443
444
Figure S7. Cluster diagram (unweighted) of all 16S rDNA and rRNA clone libraries,
445
constructed with fast UniFrac using the unweighted option, which does not take into
446
account OTU abundances. The scale bar shows the distance between clusters in UniFrac
447
units: a distance of 0 means that two samples are identical, and a distance of 0.5 means
448
that two samples contain mutually exclusive lineages. Particle-associated libraries are
449
bolded. The numbers show the support for each node as determined by jackknife
450
analysis. PA = Particle-associated; FL = Free-living.
451
Figure S8. Shannon evenness. “Max” indicates that the maximum rates of hydrolysis
452
for each substrate / incubation were used in the analysis, regardless of when the
453
maximum occurred. Sample point of the maximum rate can be seen in Fig. S2. “Day 2”
454
indicates that the hydrolysis rate for each substrate and incubation from day 2 was used in
455
the analysis.
456
16