Available online at www.sciencedirect.com
Journal of Marine Systems 74 (2008) 964 – 977
www.elsevier.com/locate/jmarsys
Microbial food web responses to light and nutrients beneath the
coastal Arctic Ocean sea ice during the winter–spring transition
Ramon Terrado a,⁎, Connie Lovejoy a , Ramon Massana b , Warwick F. Vincent c
b
a
Département de Biologie Université Laval & Québec-Ocean, Québec City, Québec, Canada G1K 7P4
Institut de Ciències del Mar CMIMA (CSIC), Dept. Biologia Marina i Oceanografia, Passeig Marítim de la Barceloneta,
37-49, 08003, Barcelona, Spain
c
Département de Biologie, Université Laval & Centre d'Études Nordiques, Québec City, Québec, Canada G1K 7P4
Received 8 February 2007; received in revised form 29 October 2007; accepted 18 November 2007
Available online 28 November 2007
Abstract
We measured the abundance and biomass of phototrophic and heterotrophic microbes in the upper mixed layer of the water column
in ice-covered Franklin Bay, Beaufort Sea, Canada, from December 2003 to May 2004, and evaluated the influence of light and
nutrients on these communities by way of a shipboard enrichment experiment. Bacterial cell concentrations showed no consistent
trends throughout the sampling period, averaging (±SD) 2.4 (0.9) × 108 cells L− 1; integrated bacterial biomass for the upper mixed
layer ranged from 1.33 mg C m− 3 to 3.60 mg C m− 3. Small cells numerically dominated the heterotrophic protist community in both
winter and spring, but in terms of biomass, protists with a diameter N 10 µm generally dominated the standing stocks. Heterotrophic
protist biomass integrated over the upper mixed layer ranged from 1.23 mg C m− 3 to 6.56 mg C m− 3. Phytoplankton biomass was low
and variable, but persisted during the winter period. The standing stock of pigment-containing protists ranged from a minimum value
of 0.38 mg C m− 3 in winter to a maximal value of 6.09 mg C m− 3 in spring and the most abundant taxa were Micromonas-like cells.
These picoprasinophytes began to increase under the ice in February and their population size was positively correlated with surface
irradiance. Despite the continuing presence of sea ice, phytoplankton biomass rose by more than an order of magnitude in the upper
mixed layer by May. The shipboard experiment in April showed that this phototrophic increase in the community was not responsive
to pulsed nutrient enrichment, with all treatments showing a strong growth response to improved irradiance conditions. Molecular
(DGGE) and microscopic analyses indicated that most components of the eukaryotic community responded positively to the light
treatment. These results show the persistence of a phototrophic inoculum throughout winter darkness, and the strong seasonal
response by arctic microbial food webs to sub-ice irradiance in early spring.
© 2007 Elsevier B.V. All rights reserved.
Keywords: Protists; Phytoplankton; Bacteria; Nutrients; Mixed layer; Sea ice; Light response
1. Introduction
Microbes are a heterogeneous group that includes
Bacteria, Archaea and small Eukarya. Given their large
⁎ Corresponding author. Fax: +1 418 656 2339.
E-mail address: [email protected] (R. Terrado).
0924-7963/$ - see front matter © 2007 Elsevier B.V. All rights reserved.
doi:10.1016/j.jmarsys.2007.11.001
standing stocks and multiple biogeochemical roles,
these communities are a key to understanding the
dynamics of marine ecosystems (Karl, 2002; DeLong
and Karl, 2005; Giovannoni and Sting, 2005; Suttle,
2005). Although bacteria and protists have been
intensively studied in the World Ocean over the last
three decades (Azam et al., 1983; DeLong and Karl,
R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
2005) there are few data from arctic waters, especially
throughout the winter season (Ducklow, 2000; Sherr and
Sherr, 2002; Calbet and Landry, 2004). Oceanographic
research in the Arctic is greatly restricted through most
of the year because of the presence of thick ice cover.
The majority of studies that have reported on microbial
dynamics in the Arctic are limited to summer months
(Thibault et al., 1999; Auel and Hagen, 2002; Hopcroft
et al., 2005; Lee and Whitledge, 2005). A notable
exception is the SHEBA/JOIS project in 1997/98 that
sampled throughout winter (Sherr and Sherr, 2003;
Sherr et al., 2003). This pioneering study showed that
microbes were active during arctic winter darkness,
although at a lower rate in comparison with summer.
Microbial communities in the Arctic are likely to be
constrained by environmental factors and the wide
fluctuation in physical and chemical conditions throughout the year. Primary production depends upon the
strong seasonality of incident irradiance, the extent and
duration of sea ice and the availability of inorganic
nutrients (Smith and Sakshaug, 1990). Bacterial metabolism is slowed by low temperatures (Pomeroy and
Deibel, 1986), while the availability of substrates for
bacterial growth is affected by autochthonous primary
production (Azam et al., 1993; Arrigo et al., 1999) and
the influx of allochthonous organic matter from rivers
(Opsahl et al., 1999; Wheeler et al., 1997), both subject
to strong seasonal variations.
The Arctic Ocean has a low-salinity, low nutrient
surface mixed layer that is separated by a strong
pycnocline from deeper, more nutrient-rich waters.
Despite this barrier to mixing, episodes of convective
exchange occur due to the formation of dense water at the
surface during freezing (Solomon, 1973; Macdonald
et al., 1989). If enough dense water is formed, it sinks
and flows off the shelf, ultimately reaching a depth where
it matches the density of the surrounding oceanic water.
As it sinks it is replaced by nutrient-rich, off-shelf waters
that advect nutrients inshore. There is considerable
interannual variation in deep water formation (Melling,
1993; Melling and Moore, 1995) and therefore in the net
nutrient influx to surface waters over the shelf.
Spring is especially important in Arctic waters. It
represents the transition from winter darkness and is a
period of rapidly increasing daylength (Burn, 1996).
Snow cover plays an important role in the penetration of
light underneath the ice, which in turn strongly affects
autotrophic production (Vincent and Belzile, 2003;
Mundy et al., 2005; Riedel et al., 2006; Mundy et al.,
2007). The transition from sea ice to open water
conditions in summer causes a major increase in
penetration of light into the water column for photo-
965
synthetic activity (Smith and Sakshaug, 1990). For
zooplankton, these changes represent the beginning of
their phase of active feeding and reproduction, which
can be strongly coupled to changes at the base of the
microbial food web (Bamstedt et al., 1999; Kosobokova, 1999; Lischka and Hagen, 2005). However, little
is known about microbial dynamics during spring, prior
to ice melt, because of the extreme difficulty of ship
access for sampling at that time of year.
The Canadian Arctic Shelf Exchange Study (CASES)
during 2003/04 provided a unique opportunity to evaluate
microbial dynamics during the period of continuous ice
cover. Our research platform (CCGS Amundsen) was
frozen into the ice in Franklin Bay, adjacent to the
Beaufort Sea, and allowed continuous sampling throughout winter and spring. Our objectives in the present study
were twofold. Firstly, we aimed to follow microbial food
web dynamics over the winter–spring period to determine
the magnitude of overwintering stocks in the arctic shelf
environment, and the timing of growth responses to the
return of sunlight. Secondly we aimed to determine the
relative importance of light exposure and nutrient supply
in controlling microbial food web dynamics during
spring. To address the latter objective we undertook a
shipboard enrichment study, and examined the responses
by microscopy and molecular methods. We applied a
pulsed versus batch set of nutrient additions using an
experimental protocol that has previously shown strong
responses in an arctic polynya.
2. Materials and methods
2.1. Sample collection
As part of the CASES program, the icebreaker CCGS
Amundsen was stationed in Franklin Bay (70° 1.3' N,
126° 25.2' W) for continuous biological and physical
studies during winter 2003 and spring 2004. This
permitted under-ice sampling of the water column
every 6 days over nearly 6 months. Samples were
collected at different depths with a rosette system
equipped with 12-L Niskin bottles and a CTD sensor
to obtain physical measurements down the water
column. During the overwintering period, the depths of
20 m and below were sampled through an opening to the
ocean from inside the ship (“moon pool”). Depths of 2 m
and 10 m were sampled with a Go-Flo (General
Oceanics) bottle from a hole on the ice 500 m away
from CCGS Amundsen, in the upstream direction of the
dominant current flow to minimize any influence from
the ship. Surface irradiance measurements were taken
from an ice camp situated 1 km from the ship.
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R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
2.2. Experimental approach
To determine the effect of nutrients on the microbial
growth, water was collected on 23 April 2003 and semicontinuous cultures were set up using a protocol that has
been previously applied to the North Water polynya
(Lovejoy et al., 2002b; Lovejoy et al., 2004). Surface
water was distributed into 6 acid cleaned 4-L Nalgene
polycarbonate bottles, after rinsing 3 times with sample
water. Initial samples for chlorophyll a (chl a), flagellates
and bacteria from the same cast were collected. Additional
surface and deep sea water were filtered through 0.2 µm
membranes to minimize biological activity in these waters
and stored in cleaned, acid-rinsed containers in the dark
and at 1 °C until utilization in the semi-continuous
cultures.
The 4-L Nalgene bottles used as microcosms were
placed in an incubator (Sanyo versatile environmental test
chamber) set at −0.4 °C in a 0 °C room in a cycle of 12 h of
dark and 12 h of light; cells were exposed to irradiance of
233 ± 25 µmol photons m− 2 s− 1 after covering with a
neutral density mesh, measured with a BSI QSL-1000
spherical PAR sensor. The bottles were pre-incubated for
2 days, after which three different treatments were applied
to duplicate bottles. The first day that the treatments were
applied was designated TS (starting time). One treatment
was designed to simulate a low nutrient supply (LOW),
and a second a high nutrient supply rate (HIGH). The
semi-continuous regime was designed to mimic in situ
sinking and grazing loss rates in the natural environment
(Hargrave et al., 2002). One liter was removed every
sampling day and replaced with comparatively low
nutrient surface 0.2 µm-filtered seawater (LOW) or with
deep nutrient-rich 0.2 µm-filtered seawater (HIGH)
(Table 1). A duplicate control treatment consisted of a
batch culture (BATCH), with no replacement of the water.
Microcosms were sampled every 2 days for chl a,
nutrients and epifluorescence cell counts; at the end of the
incubation 1 L was collected from each bottle for DNA
analysis. The subsampling time scales were chosen to
match average growth rates for polar phytoplankton
(Smith et al., 1999; Mei et al., 2002). Sampling was at the
same time of day throughout the experiment to minimize
the confounding effects of diurnal patterns, such as those
previously reported (Wheeler et al., 1989) that could
interfere with the detection of longer term trends.
2.3. Pigment and nutrient analyses
Samples for total chlorophyll a (chl a) were filtered onto
Whatman GF/F filters; the chl a for the b3 µm
(picophytoplankton) was prefiltered through a 3 µm filter
and then onto a Whatman GF/F filter. Samples were stored
frozen until they were extracted onshore in boiling ethanol
and measured fluorometrically using a Cary Eclipse
spectrofluorometer (Nusch, 1980). Water for nutrient
analysis was collected into 15 mL acid rinsed Falcon™
centrifuge tubes after prefiltration through a 0.2 µm
Sartorius Minisart syringe filter. Ammonium concentrations were estimated onboard within 24 h of collection of
samples using the method given in (Bower and HolmHansen, 1980). Nitrate+nitrite (nitrate), soluble reactive
phosphorus (SRP) and silicate (Si) were analyzed onboard
the ship, using a Brann and Luebbe autoanalyzer II and
standard protocols (Grasshoff, 1999).
2.4. Microscopy
Samples for analysis by epifluorescence microscopy
were fixed with glutaraldehyde (Electron Microscopy
Sciences) at a final concentration of 1%. Fixation took
place in the dark at 4 °C for 1 to 24 h. After fixation,
samples were filtered onto 0.2 µm (for bacteria) or 0.6 µm
(for flagellates) black 25 mm-diameter polycarbonate
Table 1
Nutrient concentrations present in the replacement waters and at
different time intervals of the incubations
Day
Nitrate
SRP
Si
HIGH replacement
LOW replacement
Ti
17.2
2.1
2.1
1.3
0.8
0.8
30.3
7.0
7.0
Ts
HIGH
LOW
BATCH
1.1 ± 0.6
1.8 ± 0.0
0.6 ± 0.9
0.5 ± 0.2
0.7 ± 0.0
0.3 ± 0.4
3.3 ± 2.0
6.6 ± 0.1
2.1 ± 2.9
Ts + 4
HIGH
LOW
BATCH
5.6 ± 2.5
1.3 ± 0.3
1.0 ± 0.1
0.7 ± 0.3
0.6 ± 0.1
0.5 ± 0.1
11.4 ± 6.0
5.0 ± 0.9
4.0 ± 0.5
Ts + 8
HIGH
LOW
BATCH
11.1 ± 0.1
1.6 ± 0.0
1.6 ± 0.1
1.1 ± 0.0
0.7 ± 0.0
0.7 ± 0.0
22.1 ± 0.1
6.8 ± 0.4
7.0 ± 0.3
Ts + 12
HIGH
LOW
BATCH
11.0 ± 0.9
0.1 ± 0.2
0.6 ± 0.5
0.9 ± 0.1
0.4 ± 0.3
0.5 ± 0.2
23.5 ± 2.4
5.0 ± 3.2
4.8 ± 3.0
Treatment
Nitrate, soluble reactive phosphorus (SRP) and silica (Si) are given in
μmol L− 1. Initial values at time of collection are given (Ti). Ts: start
value for the experiment after 2 days preincubation (average ± SD,
n = 2); Ts + 4, Ts + 8, Ts + 12: concentrations at 4 days intervals since start
of the treatments (average ± SD, n = 2).
R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
filters (Poretics) and stained with 4'6-diamino-2-phenylindole (DAPI, 5 µg mL− 1 final concentration, Sigma),
for at least 5 min. Filters were mounted onto slides with
non-fluorescent mounting oil (Zeiss Immersol 518 N).
Flagellate counts were made with a Zeiss Axiovert 100
fluorescence microscope equipped with a mercury light
source. Excitation/emission filters were 330/400 for
DAPI and 450–490/515 for chlorophyll. Total flagellate
counts were grouped into categories based on size (b2, 2–
5, 5–10 and N10 µm) and the presence/absence of
chloroplast fluorescence. The N 10 µm fraction represented cells with a diameter between 10 and 20 µm, thus
excluding identifiable ciliates, but it included diatoms
with a larger diameter in one axis. Bacteria were
enumerated following DAPI staining (Porter and Feig,
1980) either onboard the CCGS Amundsen at 1000×
magnification using an Olympus BX51 microscope or at
Université Laval with a Zeiss Axiovert 100.
Samples for inverted light microscopy were fixed
with a mixture of glutaraldehyde and buffered paraformaldehyde (1% final concentration) and stored in the
dark at 4 °C (Lovejoy et al., 2002a). With the addition of
the nucleic acid stain DAPI and use of UV and blue
fluorescent excitation filters, this technique facilitates
the differentiation of pigmented cells from heterotrophic, non-chlorophyll containing cells. After a
maximum of 8 months after collection 50 mL were
allowed to settle in sedimentation chambers for at least
24 h. Between 40% and 80% of the chamber area
(491 mm2) was examined at 400×; between 50 and 200
nanoflagellates (cells N 10 µm in diameter) per sample
were counted. Given the small volume settled, concentrations of larger rare species would be poorly estimated.
These counts were performed with a Zeiss Axiovert 100
inverted microscope at Université Laval.
2.5. DNA extraction and DGGE
During the DNA sample collection, water was
filtered through a 3 µm pore size polycarbonate filter
and the same water was then filtered through a 0.2 µm
pore size polycarbonate filter. The 3 µm and 0.2 µm
filters were conserved in separate cryovials with 2 mL of
buffer (1.8 mL of 40 mmol L− 1 EDTA; 50 mmol L− 1
Tris pH = 8.3; 0.75 mol L− 1 sucrose) at − 80 °C until
extraction. DNA extraction was carried out with a DNA
extraction kit (DNeasy Plant Mini Kit, Qiagen) following the manufacturer's instructions. The PCR amplification for the DGGE analyses was done in a BioRad
Icycler thermal cycler. The general eukaryotic primers
Euk1A and Euk516r-GC (Díez et al., 2001) were used.
PCR mixtures (25 µL) contained deoxynucleotide
967
triphosphates (dNTPs) at a concentration of 200 µM;
2 mM MgCl2; each primer at a concentration of 0.3 µM;
2.5 units of Taq DNA polymerase (New England
Biolabs); and the PCR buffer supplied with the Taq
enzyme. The PCR program and DGGE was carried out
as described in Díez et al. (2004) using a DGGE 2401Rev B model (CBS Scientific Company, Inc.).
2.6. Numerical calculations
Cell counts were converted to biovolume (Hillebrand
et al., 1999). As a general rule, all cells except diatoms
were assumed to be oblate spheres with a diameter of
1.66 µm for cells b 2 µm (as used in Booth and Horner,
(1997)), 4 µm for the fraction between 2 and 5 µm, 7 µm
diameter for the 5 to 10 µm fraction, and 15 µm diameter
for the N10 µm. For diatoms an average value obtained
after the measurement of 100 cells was used (61 µm
long and 7 µm wide). Biomass values were calculated
from biovolume using the carbon:volume relationships
in Menden-Deuer and Lessard (2000) for non-diatom
cells (C = 0.216 × V 0.939 , where C is the biomass
expressed in pg carbon and V the volume expressed in
cubic µm) and for diatoms (C = 0.288 × V0.811). Applying these relationships yields values of 0.49 pg C cell− 1
for the b 2 µm cells, 5.8 pg C cell− 1 for cells between 2
and 5 µm, 28.3 pg C cell− 1 for cells between 5 and
10 µm, 242 pg C cell− 1 for cells N10 µm (other than
diatoms), and 113 pg C cell− 1 for diatoms. These values
were comparable to those used by Sherr et al. (2003).
Bacterial biomass was estimated using a value of 10 fg
C cell− 1, a value for oligotrophic waters (see Section 4).
Biomass was integrated for the surface mixed layer
using Simpson's integration with the formula Cz1, z2 =
ΔZ × (Xz1 + Xz2 + (Xz1 × Xz2)1/2) / 3 where Cz1, z2 is the
integrated variable between depths z1 and z2, ΔZ the
distance between the two depths in m, and X is the
variable value at a given depth. These integrated values
were divided by the total depth of integration to obtain
mean concentrations.
For the experiments, in situ growth rates were
estimated from log-linear regressions of the cell counts
versus experiment day. The net change in the experiment
variables was calculated for each 2-day interval, correcting for the removal and replacement of the seawater
culture with new water. For all treatments this was:
DCt;tþ2 ¼ Ctþ2 ð1 qÞCt þ qCr
where Cr was the concentration of the replacement water
and ρ was the 2-day dilution factor. For the BATCH
control ρ was equal to 0 (no water replacement) and for
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R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
HIGH and LOW ρ was equal to 0.25. For nutrients, Cr
had a different value depending on the treatment. For the
rest of microbial variables, Cr was equal to 0 since all
components were removed by 0.2 µm filtration. All
statistical analyses were performed with SigmaStat 3.1
(Systat Software Inc.).
Table 2
Comparison of winter (December–March) and spring (April–May)
cell abundance (cells L− 1) in surface, 10 m, and inversion temperature
depth, of bacteria and size fractionated protists (average ± SD)
Depth
Abundance (cells L− 1)
Winter
Spring
Surface
10 m
Inversion temperature
2.4 ± 1.0 × 108
2.3 ± 0.7 × 108
2.1 ± 0.5 × 108
2.7 ± 1.0 × 108
2.6 ± 1.2 × 108
1.9 ± 0.7 × 108
Heterotrophic protists
b2 μm
Surface
10 m
Inversion temperature
2–5 μm
Surface
10 m
Inversion temperature
5–10 μm Surface
10 m
Inversion temperature
N10 μm
Surface
10 m
Inversion temperature
1.9 ± 0.7 × 105
2.2 ± 1.1 × 105
1.8 ± 0.8 × 105
7.2 ± 4.2 × 104
9.2 ± 4.9 × 104
7.6 ± 4.0 × 104
2.1 ± 1.1 × 104
2.8 ± 1.8 × 104
2.1 ± 1.7 × 104
7.3 ± 5.3 × 103
5.4 ± 6.8 × 103
3.4 ± 5.1 × 103
2.4 ± 1.1 × 105
2.2 ± 0.8 × 105
1.7 ± 1.3 × 105
9.5 ± 3.9 × 104
8.0 ± 3.9 × 104
5.7 ± 2.7 × 104
4.5 ± 1.0 × 104
3.4 ± 1.5 × 104
1.3 ± 0.7 × 104
1.7 ± 1.0 × 104
1.6 ± 1.0 × 104
6.6 ± 5.3 × 103
Autotrophic protists
b2 μm
Surface
10 m
Inversion temperature
2–5 μm
Surface
10 m
Inversion temperature
5–10 μm Surface
10 m
Inversion temperature
N10 μm
Surface
10 m
Inversion temperature
7.0 ± 4.7 × 104
7.1 ± 4.8 × 104
5.0 ± 6.3 × 104
1.9 ± 1.8 × 104
1.9 ± 0.8 × 104
1.7 ± 2.2 × 104
1.1 ± 0.9 × 104
3.8 ± 4.0 × 103
7.3 ± 8.1 × 103
3.9 ± 4.7 × 103
3.6 ± 3.7 × 103
2.7 ± 5.3 × 103
2.9 ± 1.6 × 105
1.8 ± 1.8 × 105
2.4 ± 1.2 × 104
1.9 ± 0.8 × 105
1.2 ± 0.8 × 105
2.7 ± 2.6 × 103
4.0 ± 3.3 × 104
3.0 ± 3.4 × 104
1.6 ± 2.4 × 103
1.5 ± 0.8 × 104
1.6 ± 1.0 × 104
2.0 ± 2.1 × 103
Bacteria
3. Results
3.1. Franklin Bay: physical and biological parameters
Throughout the period of sampling in winter and
spring, the water column was ice-covered and salinity
stratified with a characteristic halocline. A salinity of
31.1 was used as an indicator of the upper limit of this
halocline, and over winter and spring this occurred at an
average depth of circa 25 m. The bottom limit of the
cold-water mass below the surface halocline was at a
depth of circa 125 m. Below this was a layer of water
with salinity greater than 33 and relatively warmer
temperatures (ranging − 1.25° to 0 °C). This layer
continued to the maximum water column depth of
200 m. During winter and spring a temperature
inversion could be observed due to the presence of
warmer water between two cold-water masses in the
surface layer, at depth 32.4 ± 7.3 m (average ± SD). This
temperature inversion was associated with the halocline
so it was considered as a reliable index of the depth of
the surface mixed layer.
Nutrient concentrations remained constant throughout winter and spring. Nitrate concentrations in the
upper mixed layer were usually b 3 μmol L− 1 and SRP
concentrations b 1 μmol L− 1; maximum silicate concentrations were ca. 10 μmol L− 1 in winter, with lower
values in spring (circa 7 μmol L− 1). Waters below the
halocline showed higher nutrient concentrations,
achieving maximum values of 17 μmol L− 1 for nitrate,
1.7 for SRP, and 33 μmol L− 1 for silica (data provided
by J-E Tremblay, NM Price and K. Simpson, McGill
University, Québec, Canada).
Bacterial concentrations (Table 2) were similar
between winter and spring (demarcated as before and
after March 21). The bacterial population was fairly
homogeneous down the water column and average
(± SD) concentrations were 2.4 (0.9) × 108 cells L− 1 in
the upper mixed layer.
Eukaryotic microbes were present throughout the
water column but concentrations were greater in the
upper mixed layer. The most abundant size fraction for
both phototrophic and heterotrophic organisms was the
b2 µm size category (Table 2). Although the exact
identification of phytoplankton is not possible under
epifluorescence microscopy, we noted that the b2 µm
phototrophs mostly corresponded to the characteristic
Micromonas-like cell form, with one acentric chloroplast. For the photosynthetic fraction of 2 to 5 µm, a
Phaeocystis-like cell form was the dominant, with two
chloroplasts on either side of the nucleus. In the larger
fraction (N 10 µm diameter), dinoflagellate cysts were
observed during winter and spring in the surface waters,
and in winter they were the dominant cells in this size
fraction. Diatoms in winter represented roughly 20% of
N10 µm cells, but this increased to 30% in April and
60% in early May.
Estimates of chl a concentrations generally followed
the same patterns as phytoplankton cell densities.
Surface chl a increased as winter progressed into spring
(Fig. 1). Size fractionation analyses showed that the
b3 µm fraction represented on average (± SD) 60 (15) %
R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
of the total chl a in surface and 64 (18) % of the total chl
a at 10 m. For the inversion temperature depth a lower
proportion of chl a was in the b3 μm fraction: 39
(24) %. Chl a was positively correlated with PAR
(Pearson correlation, r = 0.778, p b 0.01) and negatively
correlated with nitrate (Pearson correlation, r = − 0.771,
p b 0.01) (Fig. 1).
The average bacterial biomass concentration from the
surface to the inversion temperature depth ranged from
1.33 mg C m− 3 to 3.60 mg C m− 3 (Fig. 2) with no
significant changes between winter and spring
(ANOVA, F = 0.0143, df = 21, p N 0.05). Mean planktonic heterotrophic protist biomass for the same layer
(Fig. 2) ranged from 1.23 mg C m− 3 to 6.56 mg C m− 3
(Table 3), with no significant difference between winter
and spring (ANOVA, F = 1.797, df = 18, p N 0.05).
Picoflagellates were the most abundant eukaryotic
group by cell numbers. However in terms of biomass,
they represented 3.7 ± 2.5% (average ± SD) of the total
integrated protist heterotrophic biomass.
Phototrophic protist biomass concentrations (Fig. 2)
showed significant differences between winter and
spring (ANOVA, F = 11.193, df = 18, p b 0.01), and
ranged from a minimum value of 0.38 mg C m− 3 in
winter to a maximal value of 6.09 mg C m− 3 in spring
(Table 3). As with the heterotrophic protists, the
picophytoflagellates were generally the most abundant
in terms of cell numbers but in terms of biomass
represented only 5.1 ± 3.2% (average ± SD) in winter and
2.5 ± 1.0% in spring of the total integrated phototrophic
biomass.
969
Fig. 2. Average heterotrophic protist, autotrophic protist and bacterial
biomass at the overwintering station 200 (mg C m− 3) for the surface
mixed layer.
3.2. Nutrient influence on the development of a spring
community: experimental results
Nutrient concentrations in the surface water used to
start the incubations were moderately low (Table 1).
Ammonium was below the detection limit and remained
undetectable throughout the entire incubation. Statistical
analysis of the accumulated consumption of nutrients
during the incubations showed no significant differences
among the three different treatments for nitrate, SRP and
silica (Two way ANOVA, Nitrate: F = 8.515, df = 35,
p N 0.05; SRP: F = 3.193, df = 35, p N 0.05; Silica:
F = 3,258, df = 35, p N 0.05). Bacterial concentrations
Table 3
Comparison of winter (December–March) and spring (April–May)
average mixed layer biomass (mg C m− 3) of bacteria and size
fractionated protists (average ± SD)
Biomass (mg C m− 3)
Fig. 1. Surface chl a concentrations, incident PAR (daily integrals) and
nitrate concentrations in the surface layer during winter and spring in
Franklin Bay.
Winter
Spring
Bacteria
Total protists
Heterotrophic protists
b2 μm
2–5 μm
5–10 μm
N10 μm
2.1 ± 0.5
3.3 ± 1.6
2.5 ± 1.4
0.08 ± 0.03
0.4 ± 0.1
0.6 ± 0.3
1.4 ± 1.1
2.3 ± 0.8
6.2 ± 3.4
3.5 ± 1.9
0.08 ± 0.02
0.4 ± 0.2
0.7 ± 0.3
2.4 ± 1.6
Autotrophic protists
b2 μm
2–5 μm
5–10 μm
N10 μm
0.8 ± 0.7
0.03 ± 0.03
0.1 ± 0.08
0.1 ± 0.09
0.5 ± 0.5
2.6 ± 1.8
0.06 ± 0.02
0.4 ± 0.3
0.6 ± 0.7
1.6 ± 0.9
970
R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
were initially 2.02 × 10 5 cells mL − 1 and average (± SD)
final concentrations were similar in all treatments
(3.80 ± 1.16 × 10 5 cells mL − 1 ).
Heterotrophic flagellate concentrations and net
biomass production increased in all three treatments
(Fig. 3). The greatest value for net heterotrophic
production was in the LOW treatment, followed by
the HIGH and the BATCH, but these differences were
not statistically significant (Two way ANOVA,
F = 1.461, df = 35, p N 0.05). The last day of the
incubation had the highest concentrations of chl a, and
in both LOW and HIGH treatments the greatest increase
was from TS + 8 to TS + 10 (no pigment data were available
for the BATCH treatment because of the limited sample
volume). The concentration of phototrophic protists
increased in all three treatments, with cell concentrations
increasing an order of magnitude from that measured at
Ti. For net phototrophic biomass (Fig. 3), LOW
achieved the greatest values, followed by the BATCH
and the HIGH; however, these differences were not
statistically significant (Two way ANOVA, F = 2.310,
df = 35, p N 0.05). The b 2 µm cell fraction mostly
consisted of Micromonas-like cells and the fraction
between 2 and 5 µm was mostly Phaeocystis-like cells.
Diatoms were not abundant at the start of the experiment
(20% of the N 10 μm fraction) however between TS + 10
and TS + 12 they increased markedly and accounted for
90% of the protists in the N 10 μm fraction. Nitzschia
spp. colonies were the most abundant for all the three
treatments, with Fragilariopsis spp., Navicula spp. and
Skeletonema spp. also present.
Net in situ growth rates were estimated for the
different size fractions. Doubling times (calculated as ln
2 / µ) for heterotrophic flagellates were 18 days (b2 µm
fraction), 15 days (2–5 µm fraction) and 11 days
(N10 µm fraction). Phototrophic flagellates presented
doubling times of 10 days (for both the b 2 µm and 2–
5 µm fractions) and 7 days (N10 µm fraction). A growth
rate for the 5–10 µm fractions could not be calculated
because of the variable responses through time.
All incubations had similar biomass proportions
among phototrophs and heterotrophs at the end of the
incubation (Fig. 4). Also, the genetic fingerprint of the
final community as assessed by DGGE was similar in all
the incubations (Fig. 5). The b 3 µm fraction showed
some minor differences; however, the major bands were
present in all samples. The N3 µm fraction showed the
exact band composition among treatments, confirming
the microscope observations that all treatments had
similar final communities.
4. Discussion
Fig. 3. Average (± SE) net cumulative biomass production of
heterotrophic protists (μmol C L−1), from Ts to Ts + 12, B: Average
(±SE) net cumulative biomass production of photosynthetic protists
(μmol C L−1) from Ts to Ts + 12.
Microbial biomass at all sampling times in the
Franklin Bay shelf site was low compared with other
studies in the Arctic and elsewhere (Table 4). In part this
is related to cell concentrations. Low cell numbers were
found throughout winter and spring, resulting in mean
and integrated biomass values that are at the lower end
of the range previously reported for polar oceans.
However, another factor that affects biomass estimates
is the selection of an appropriate cell number to biomass
conversion factor, especially for bacteria. The exact
carbon content of marine bacteria is still uncertain. A
range of different conversion factors have been
R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
971
Fig. 4. Biomass size structure of heterotrophic and autotrophic protist community. Initial values at time of collection are given (Ti) and average values
for each of the treatments at Ts + 12. HP: heterotrophic protist; AP: autotrophic protist.
proposed that relate either cell numbers or biovolume to
biomass (Watson et al., 1977; Bratbak and Dundas,
1984; Bratbak, 1985; Nagata, 1986; Lee and Fuhrman,
1987; Borsheim et al., 1990; Psenner, 1990; Fukuda
et al., 1998; Ducklow, 2000). In arctic waters a factor of
20 fg C cell− 1 factor has been used (Sherr et al., 1997,
2003; Sherr and Sherr, 2003), however given the
oligotrophic nature of this polar ocean, lower values
are likely to be more appropriate. For a low conversion
factor of 120 fg C µm− 3 (Gasol et al., 1997) an average
value of 10 fg C cell− 1 is obtained using the biovolume
data for the Arctic Ocean presented by Sherr et al.
(1997), 13 fg C cell− 1 for bacterial biovolume data of
Disko Bay (Nielsen and Hansen, 1995) and 7 fg C cell− 1
for the Antarctic Peninsula (Vaqué et al., 2002). In the
present study in the Beaufort Sea, considering the small
cell characteristics of the bacteria observed and the
oligotrophic nature of the sampled zone, a value of 10 fg
C cell− 1 was considered most appropriate to convert
bacterial counts to bacterial biomass instead of the
widely used value of 20 fg C cell− 1. In the case of
protists, an average value was considered for each size
group counted under the microscope, which gave
biomass conversion factors comparable to those found
in the literature for the Arctic Ocean (Sherr et al., 2003).
Heterotrophic bacteria and flagellates mostly dominated winter microbial biomass. Bacteria were active
through winter and production peaked in summer
(Garneau et al., in press). Ciliates were also present
during winter and spring, although their biomass was
lower than those of heterotrophic flagellates (Vaqué
et al., submitted for publication). Our microscope
observations revealed the persistence of pigmented
cells throughout winter despite the extremely low
irradiance levels caused by the snow and ice cover in
combination with low incident irradiance. The chl a and
cell concentrations were small (Fig. 1; Table 2), and the
dominant fraction of this phototrophic community
972
R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
Fig. 5. Negative images of DGGE gel of the N3 µm and the b3 µm assemblage at the end of the incubation of the different microcosms.
corresponded to Micromonas- like cells, consistent with
HPLC pigment analyses of the community structure
(Lovejoy et al., 2007). Larger Phaeocystis-like cells and
diatoms were also common. Most of the diatoms
sampled at this time of year did not appear to be healthy
and their chlorophyll appeared degraded under the
fluorescence microscope. In contrast, the chlorophyll in
the Micromonas-like cells was brightly fluorescent. It is
interesting that a positive environmental growth rate
from February to April was found for this population
(Lovejoy et al., 2007) indicating that they retained
viability over winter and an ability to respond
immediately to improved light conditions. Sherr et al.
(2003) suggested that Micromonas sp. might be capable
Table 4
Average biomass values (mg C m− 3) of bacteria, heterotrophic protists and autotrophic protists (b20 µm) for different oceanic regions
Region
Equatorial Pacific Ocean
North Atlantic
Sub arctic Pacific
Disko Bay (Greenland)
Antarctic Peninsula
Arctic Ocean
Arctic Ocean
Arctic Ocean
Beaufort Sea
Beaufort Sea
Month–year
July 1998
May 1989
Feb 1994, May 1994/95
June–July 1992
November–February 1995/96
July–September 1994
November 1997–June 1998
June–August 1998
December 2003–March 2004
March 2004–May 2004
Average biomass (mg C m−3)
Bacteria Heterotrophic protists Autotrophic Integration
b20 μm
protists
Reference
–
36.9
–
53.5
6.1
9.7
3.5
6.4
4.2 (2.1)
4.6 (2.3)
(Yang et al., 2004)
(Sieracki et al., 1993)
(Boyd and Harrison 1999)
(Nielsen and Hansen 1995)
(Vaqué et al., 2002)
(Sherr et al., 1997)
(Sherr et al., 2003)
(Sherr et al., 2003)
This study.
This study.
6.8
30.4
–
11.5
0.7
6.6
1.9
4.5
2.5
3.5
19.0
58.3
4.9
83
2.9
40.5
0.4
10.9
0.8
2.6
Mixed layer
30 m
Mixed layer
Photic zone
40 m
50 m
40 m
40 m
Mixed layer
Mixed layer
Values were obtained from the literature directly or transformed to present the integrated values. Transformations from chl a values were done using
C:chl a average ratio from (Booth and Horner 1997). Transformations to bacterial biomass from bacteria concentrations were done using the
conversion value of 20 fg C per cell (Lee and Fuhrman 1987) and for this study the values using a conversion factor of 10 fg C per cell is also
presented in parenthesis (see Section 4 for details).
R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
of phagotrophy, which could be a mechanism contributing to their winter survival, reported for some phototrophic flagellates in polar lakes (Bell and LaybournParry, 2003; Laybourn-Parry et al., 2005). Cells
identified as Micromonas pusilla are reported to be
capable of phagotrophy elsewhere (González et al.,
1993). We have no direct evidence from the present
study, since no Micromonas-like cells were recorded
with DAPI-stained fluorescent particles inside, and this
possibility requires further evaluation.
Winter community biomass was dominated by
heterotrophic protists, and as spring advanced, phototrophic organisms represented an increasing proportion
of the total biomass (Fig. 2). Chl a was positively
correlated with PAR, implying a growth response to
irradiance. The availability of this light for phytoplankton growth may also have been influenced by the snow
melt from the ice cover, as noted by Sherr et al. (2003).
Chl a was negatively correlated with nitrate concentrations, implying active growth and nutrient depletion.
Although nitrate is often thought of as a substrate for
larger phytoplankton rather than small cells, recent
genomic analyses of Ostreococcus, another picoprasinophyte closely related to Micromonas, have shown
genes coding for transport systems for nitrate in addition
to urea and ammonium (Derelle et al., 2006). Parallel to
the increase in phototrophic biomass there was a change
in community structure. With increasing irradiance
during spring, an increase in diatom biomass was
observed, although they did not reach bloom concentrations. Diatoms in spring represented as much as 60% of
the N 10 μm cells while in winter they only represented
20% and microscope observations showed these cells
were mostly degraded. The carbon estimates for large
phytoplankton are equivalent to the pigment values
presented in Lovejoy et al. (2007). On the other hand,
our values of carbon biomass for small-celled phytoplankton are lower. If a ratio carbon:chl a is calculated
for the N3 µm fraction, the average ratio is 40. For the
b 3 µm, this ratio falls to 10. This suggests that our
carbon values for the picophytoplankton may be
underestimated, which may be an artifact of our
microscope counts. The signal from DAPI stained
cells when observed under the microscope is likely to
fade more rapidly for small cells than large cells.
The pycnocline was well established throughout the
period of sampling and effectively sealed the nutrients in
the deep water, preventing replenishment of the surface
mixed layer. Unlike other environments, there was no
spring bloom and chl a never rose beyond 1 μg L− 1 in
Franklin Bay. Nutrient concentrations in surface waters
were low throughout winter, and our enrichment
973
incubations provided an experimental test of whether
nutrient supply was a limiting factor during the initial
phase of the winter–spring transition. The water for
these incubations was collected from the upper water
column covered by 1.8 m of sea ice, and the experiment
simulated a fast retreat of the ice with exposure to
irradiance levels similar to those found at surface open
waters at that time of the year. The two treatments with
water replacement were designed to simulate different
nutrient flux conditions. The LOW treatment simulated
low nutrient conditions for the upper mixed layer and at
the top of the halocline. The HIGH treatment simulated
the entrainment of nutrients into the upper mixed layer
from below. The N:P ratio of dissolved nutrients in the
HIGH replacement waters (13.1:1) was close to the
Redfield ratio of 16:1, a common feature of deep waters,
while the LOW replacement waters had a lower ratio
(2.6:1) indicating that these waters were potentially
more limiting in nitrate compared to phosphorus
(Guildford and Hecky, 2000). The N:Si ratio of both
replacement waters was below 1, suggesting again that
nitrate would be the nutrient most likely to be limiting
(Levasseur and Therriault, 1987). However, although
nitrate was always potentially limiting, it was never
depleted over the course of the experiments, even in the
BATCH control culture (Table 1), suggesting that
phytoplankton were never strongly nutrient-limited.
There were no differences in the total consumption of
nutrients among treatments. Even in the case where
more nutrients were added in the HIGH treatment, the
microbial community did not respond to this enrichment. The ammonium levels were always below the
limit of detection, indicating that any production was
consumed as fast as it was produced, consistent with
ammonium as the preferred source of nitrogen (Harrison
et al., 1996).
Despite the different nutrient conditions, no differences were detected among treatments in either biomass
production or in community structure. A marked shift in
community composition was evident at the end of the
experiment relative to starting conditions; however, the
final proportions of the different size fractions were
similar among treatments (Fig. 4). The initial community
at TS was dominated by heterotrophs (N50% of total
biomass) and by the end of the experiment, at TS + 12, all
of the microcosms were dominated by phototrophs,
which contributed N 80% of the total protist biomass. At
TS the phototrophic community biomass was dominated
by small organisms (b 10 µm), and diatoms represented
only a small percentage (ca. 20%) of the N 10 µm
fraction. At the end of the experiment the N10 µm
fraction contributed a greater proportion of biomass, and
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R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
was composed mostly of diatoms. This shift towards a
phototrophic, diatom-rich community was not influenced by nutrient conditions.
Other factors could have influenced phytoplankton
growth during the course of our incubations. For
example, vitamin B12 is an important micronutrient
for large phytoplankton (Sañudo-Wilhelmy et al., 2006),
and it has been suggested that bacteria may be an
important source of this vitamin for phytoplankton
(Croft et al., 2005). In our incubations, bacterial biomass
increased towards the end of the experiment and may
have resulted in increased vitamin B12 availability, or
may have had other stimulatory effects on phytoplankton growth.
A genetic fingerprinting analysis, DGGE, confirmed
that the different treatments gave rise to similar protist
communities in all the incubations. The band patterns
that defined the community composition at the end of
the incubations (Fig. 5) did not show major changes
among the treatments, with only minor shifts in the
b3 µm fraction and undetectable change in N 3 µm.
Excision and sequencing of the most prominent bands
was attempted, but the majority of sequenced bands
were of poor quality or double sequences. The DGGE
technique is largely qualitative and clean separation of
phylotypes depends mostly on relative GC content.
Given that similar GC contents are found in many small
eukaryotes in Arctic waters (C. Lovejoy & M. Potvin
unpublished data), it was impossible to assign a single
band to a single species. Despite this, the strong
similarity of band patterns among the treatments
suggested that the initial winter community had
moved towards a new species structure once exposed
to an improved irradiance regime, and that this effect
occurred irrespective of differences in nutrient supply.
Rose and Caron (2007) have suggested that in terms
of growth rates, heterotrophic protists in cold waters are
more affected by low temperature than autotrophic
protists. They suggested that this might assist the
development of phytoplankton blooms particularly in
the Polar Regions where there is already a mismatch
between early diatom production and calanoid copepod
abundance. In our incubations the growth rates of
heterotrophic flagellates were lower than those of
autotrophs, consistent with this notion.
Our experimental results contrast with those using a
similar protocol in early June 1998 in the North Water
Polynya (Lovejoy et al., 2004). The major finding of
those experiments was that advective nutrient input
resulted in the production of a diatom bloom. In our case
we did not induce a diatom bloom, although the
proportion of diatoms at the end of the incubations
was much higher than at the beginning. Heterotrophic
flagellates, which were typical of the winter community,
dominated the starting community of our incubations.
Exposing this community to increased irradiance levels
promoted the development of photosynthetic organisms.
At the end of the incubations, independent of nutrient
conditions, the community was dominated by phototrophs, but not in the form of a diatom bloom. In the case
of the Lovejoy et al. (2004) study, the initial community
was already an established spring community, with a
high proportion of photosynthetic organisms, and the
input of nutrients caused a successional change in
community structure and biomass. This was not the case
in our microbial community, where the transition from a
winter to a spring community was not driven by
nutrients but by light availability. Similarly, Sherr
et al. (2003) found an increase of Micromonas sp. and
large phytoplankton under the ice cover in spring during
the SHEBA project and attributed this growth to
increasing day length and the melting snow cover.
These two factors would also contribute to the increase
in phototrophic cells in Franklin Bay in spring. In
contrast, the influence of increasing nutrients may be
restricted to open water conditions (Sieracki et al., 1993;
Boyd and Harrison, 1999; Klein et al., 2002; Yang et al.,
2004).
The winter–spring transition is of special importance
to higher trophic levels. During spring there was an
increase in both phototrophic and heterotrophic biomass
(Table 3), which can be a food resource for herbivorous
and omnivorous copepods in the Arctic basin (Thibault
et al., 1999). The increase of protist biomass was most
pronounced for the N 10 μm cells, which are the
preferred prey of zooplankton (Stevens et al., 2004).
Copepods may survive winter on lipid reserves without
active feeding (Scott et al., 2000) but in spring, their
feeding and reproductive strategies are strongly coupled
to primary production (Bamstedt et al., 1999; Kosobokova, 1999; Lischka and Hagen, 2005). The timing and
changes in microbial community structure during the
winter–spring transition are therefore likely to have a
direct influence on the transfer of organic matter and
energy to higher trophic levels.
5. Conclusion
Our results show that a microbial community persists
throughout winter in the Arctic shelf environment.
Heterotrophic organisms dominated the winter community, but a small community of phototrophs was also
maintained throughout winter. The relative abundance
of phototrophs increased rapidly during the winter–
R. Terrado et al. / Journal of Marine Systems 74 (2008) 964–977
spring transition, which would have implications for
higher trophic levels. Our experimental study indicated
that this transition from a heterotrophic winter community to a phototrophic spring community was little
influenced by nutrient supply. Our experiments under
different nutrient regimes did not show major differences in the community dynamics, which in all cases
moved from a heterotrophic dominated community to an
phototrophic dominated community in response to
increased irradiance. These results are consistent with
the observations made during a previous study of an
Arctic winter–spring transition (Sherr et al., 2003) and
imply a general response under arctic sea ice to
improved irradiance conditions during the winter–
spring transition, with little control by nutrient supply
at that time of year.
Acknowledgements
Financial support for this study was provided by the
Natural Sciences and Engineering Research Council of
Canada, the Canada Research Chair program, Fonds
québécois de recherche sur la nature et les technologies
and Indian and Northern Affairs Canada. We thank the
officers and crew of the CCGS Amundsen, and C.
Martineau, S. Roy, M. Rautio, K. Simpson, J-E.
Tremblay, N.M. Price, A. Riedel, M. P. Lizotte and T.
Papakyriakou for field assistance and analysis. This
work was conducted within the Canadian Arctic Shelf
Exchange Study (CASES) led by L. Fortier.
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