ELSEVIER
FEMS Microbiology Ecology 16 (1995) 213-222
Hydrolytic enzymatic activity in deep-sea sediments
Knut Poremba
Forschungs- and Technologiezentrum
We&&e,
*
Universitiit Kiel, D-25761 Biisum, Germany
Received 23 March 1994; revised 1 November 1994; accepted 5 November 1994
Abstract
Hydrolytic activities of five enzymes were measured in deep-sea sediment cores at three stations under in situ
temperature and pressure in the NE-Atlantic in March/April
and July/August
1992. Generally, activity profiles declined
vertically in the upper 10 cm of the cores. Experiments under in situ pressure were not significantly different from
measurements
under surface conditions. The ranking of potential activity rates in the top sediment horizon was:
aminopeptidase > esterase > chitobiase > P-glucosidase > cr-glucosidase with ratios of 687/174/11/3/l.
This is similar to
ratios obtained in marine aggregates from the upper mixed layer, thus supporting the idea of pelagic-benthic coupling in the
open ocean. The vertical activity profiles show that the biochemical composition, and thereby the nutritive quality of the
degradable material, changed with depth in the sediment cores. About 518 mg carbon was potentially mobilized in the O-l
cm sediment horizon per square meter per day. This contrasts with the input of particulate organic carbon to the sea floor in
this area of only 2.74 mg C m2 d- ‘, determined by sediment traps, which indicates that the deep-sea benthic community can
rapidly utilize sedimenting particulate organic material and highlights the importance of extracellular enzyme activity in the
sediment biogeochemical loop.
Keywords:
Microbial activity; Enzyme activity; Deep sea; Benthos; Pelagic-benthic
1. Introduction
Particulate organic matter (POM) is an important
nutrient source for life at the deep-sea floor. Since
the benthic biomass consists mostly (> 90%) of
bacteria [II, which can incorporate only low-molecular-mass
compounds,
the extracellular
enzymatic
cleavage of POM into small molecules is a key step
in benthic metabolism [2-41. Extracellular
enzyme
activity is usually measured with fluorogenic substrate analogs, focusing primarily on aminopepidases
* Corresponding author. Tel.: + 49 (4834) 604206. Fax: + 49
(4831) 604299.
0168-6496/95/$09.50
SSDIO168-6496(94)00085-9
coupling
and glycosidases [4]. An additional commonly used
model substrate is fluorescein diacetate (FDA), representing an esterase activity although this is not
exclusively extracellular 151.
Despite the ocean bottom clearly being a site of
intense biological
activity and of importance
for
marine nutrient cycles, most investigations on hydrolytic activity in the open sea have been carried out in
the water column [6-91. Due to logistic and financial
problems, measurements on hydrolytic activity in the
deep-sea have been conducted
only sporadically
[lo-121.
The most striking factor distinguishing
the deepsea from other environments
is the elevated hydrostatic pressure, so the proposition that deep-sea bac-
0 1995 Federation of European Microbiological Societies. All rights
reserved
214
K. Poremba /FEMS
Microbiology
Ecology 16 (19951213-222
teria are physiologically
adapted to pressure, is old
[13]. Several investigations
had clearly shown the
existence of microbial consortia in the deep-sea,
which show higher activity under their in situ pressure [14-161. By using an improved water sampler
in the Mediterranean
Sea, barophilic microrganisms
have been found in a shallow water depth of 1100 m
[17,18]. Therefore, the use of in situ measurement
systems or the performance
of ship-board incubations under elevated pressure are recommended
for
the measurement
of biological activity in the deep
layers of the ocean.
The aim of the investigation reported here was to
measure (i) the patterns of five hydrolytic enzyme
activities (aminopeptidase,
esterase, chitobiase,
CYand /3-glucosidase) in deep-sea sediment, (ii> their
spatial variation in the NE-Atlantic
on a transect
along 19” W and between 47” and 59” N, (iii> their
temporal variation between spring and summer 1992,
and (iv> the effect of pressure on activity.
2. Material and methods
2.1. Sampling strategy and ship-board
handling
To study the patterns and spatial variability of
enzyme activity in sediment of the NE-Atlantic,
samples were taken in the West European Basin and
in the Iceland Basin. Temporal variability was examined by comparing esterase rates in the West European Basin during spring and summer.
Sediment samples were collected with a multiple
corer [19] on two cruises of the R.V. Meteor
(March/April
and July/August
1992) in the NEAtlantic (Fig. 1, Table 1). The samples were pro-
Table 1
Sampling
60’
60’
40’
ZO’W
0’
Fig. 1. Map of sampling sites.
cessed immediately for measurements of various hydrolytic activities. The assays were performed in an
experimental container, in which in situ temperature
and pressure were simulated.
2.2. Determination
of bacterial number
The total bacterial number (TBN) was determined
according to a modified version of the method of
Hobbie et al. [20]. Sub-cores (length: 15 cm; diameter: 1.39 cm) were taken from one multiple corer
tube and sectioned into 1 cm horizons. Each horizon
was diluted immediately up to tenfold in pre-filtered
(0.2 pm) sea water supplemented
with formaldehyde (4% final concentration)
and stored cool and
sites
Station number
20’E
Date
Location
21th March 1992
27th March 1992
28th March 1992
1st April 1992
28th July 1992
11th August 1992
15th August 1992
22th August 1992
46O23.85’
47O10.74’
47”10.87’
47YO.86’
59”16.23’
47”10.76’
47O10.77’
47YO.86
Depth Cm)
N
N
N
N
N
N
N
N
d35.58’ W
19O33.48 W
19O33.57’ W
19O33.73’ W
21”04.80 W
19O33.77’ W
19O33.25’ W
19”33.76’ W
4919
4568
4569
4575
2879
4564
4565
4565
K. Poremba /FEMS
Microbiology
dark for later analysis. In the laboratory the samples
were diluted up to lOO-fold, ultrasonicated
for 5 s,
and diluted further up to lOOOO-fold. An appropriate
volume of 2-5 ml was filtered onto a 0.2 pm
Nuclepore filter, stained for 2 min with 0.1% acridine orange, and washed with citrate buffer (0.056 M
Na-citrate, 0.056 M NaOH, 0.044 M HCl, pH 4).
Stained bacteria were counted with an epifluorescence microscope.
2.3. Assay for hydrolytic
activities
Measurements
of enzyme activities were performed by adding fluorescent substrate analogs to 10
ml sediment slurries. The model substrates (all purchased from Serva Feinbiochemica,
Heidelberg) are
shown in Table 2. Their concentrations
used during
the experiments corresponded with substrate saturation levels of the enzymes, which had been previously determined. The drawback of preparing slurries is that the original structure of the sediment is
destroyed. Injection of substrate directly into intact
sediment cores [21,22] can avoid this problem but
leads to further difficulties due to different diffusion
rates and an unequal distribution of the substrate. To
minimize this drawback of the slurry technique, the
effects of increasing (l:l, 1:3, 1:9, 1:19) sediment
dilutions were tested with sediment from station 6.
All other experiments were performed by using 1:l
sediment dilutions.
The sediment slurries were prepared by sectioning
1 cm horizons of three cores, pooling them, and
diluting them 1:l with sterilized deep water. The
slurries were pipetted in plastic bags, supplemented
with fluorescent substrates and incubated at in situ
temperature. One set of bags was kept at 1 atm while
the other set was put into pressure vessels, in which
in situ pressure conditions were produced using an
Table 2
Model substrates
Type of enzymatic
Aminopeptidase
Esterase
Chitobiase
P-Glucosidase
a-Glucosidase
used for measuring
reaction
hydrolytic
Ecology 16 (1995) 213-222
21.5
electronic hydrologic pump (pressurization
and depressurization:
about 100 atm min- ’ ). The experiments were terminated after 0, 2, 4, or 6 h, after
which 1 ml of the slurry was diluted with ice-cold 9
ml sea-water and centrifuged at 5000 X g. Series
with autoclaved samples served as controls. Fluorescence was determined with a Jasco FP-550 spectrofluorometer
calibrated with known amounts of
7-amino-4-methylcoumarin,
fluorescein, and 4-methylumbelliferone.
Assays were done in triplicate. Assuming that 1 mol enzymatically cleaved fluorogenic
substrate corresponds to the liberation of 4 mol C (2
mol acetate), or 6 mol C (leucine, glucoside, glucosamine) the activities of aminopeptidase,
esterase,
chitobiase, /3- and cr-glucosidase were calculated as
mg C potentially mobilized via hydrolytic cleavage
per square meter of the top cm of the sediment per
day. The ratios of enzyme activities were calculated
with reference to the a-glucosidase activity.
3. Results
Assays of enzymatic activity were performed using sediment slurries, which were supplemented with
fluorescent substrate analogs. To test possible effects
of increasing dilutions were tested, cleavage rates
obtained from series with up to 20 X sediment dilution were compared (Table 3). The results indicated
that the assay leads to an increasing underestimation
of activity in the top sediment with each additional
dilution step. In deeper layers, this effect of dilution
was negligible. Therefore, a limited dilution of only
1:l (v/v> was used subsequently in all our experiments.
Activity profiles of the five hydrolytic enzymes
are presented in Tables 4 and 5. In general the
highest rates were found in the top horizons decreas-
enzyme activity
Substrate analog
Substrate concentration
used(pmoll-t)
L-leucine-7-amino-4-methylcoumarin
fluorescein diacetate
4-methylumbelliferyl-N-acetyl-/3-pglucosamide
4-methylumbelliferyl-P_Dglucopyranoside
4-methylumbelliferyl-cY_Dglucopyranoside
1000
100
100
100
50
K. Poremba / FEMS Microbiology
216
Table 3
Effect of diluting deep-sea sediment
diacetate (FDA) cleavage rates a
Horizon
km>
O-1
3-4
7-8
(station
6) on fluorescein
Sediment
dilution
FDA hydrolysis rate
(nmol cmm3 h-r )
Correlation
(r?
1:l
1:3
1:9
1:19
1:l
1:3
1:9
1:19
1:l
1:3
1:9
1:19
7.264
4.303
3.180
1.585
3.307
3.718
2.553
3.384
3.442
2.699
2.271
1.363
0.997
0.989
0.992
0.971
0.983
0.982
0.976
0.946
0.975
0.978
0.973
0.961
a Pressure during incubations
= 1 atm.
ing with depth into the sediment.
However,
aminopeptidase
always showed a small intermediate
maximum at 2-4 cm depth.
The shape of the activity profile was dependent
on the enzyme. In the upper 10 cm of the sediment,
aminopeptidase
activity decreased less (to 39% of
the surface value) than the esterase (to S%,>, while
and
glucosidase
decreased
most
chitobiase
(chitobiase: to 3%; both glucosidases: to almost 0%)
down to the detection limit of the method (about
0.002 nmol cme3 h- ‘).
Table 4
Hydrolytic
activities
a (nmol cm -3 h- ‘) in deep-sea sediments
Sediment depth (cm)
Aminopeptidase
o-1
l-2
2-3
3-4
4-6
6-8
8-10
Sediment depth (cm)
22.351 +
19.482 +
22.511 +
23.029 +
19.141 f
16.221 f
10.044 f
Chitobiase
O-1
0.285
0.177
0.118
0.089
0.065
0.046
0.007
l-2
2-3
3-4
4-6
6-8
8-10
a All values are means f standard
k
f
+
f
+
f
+
(stn. 2, 3, 5, 7, 8)
5.041
3.044
3.842
5.417
6.462
6.818
3.944
(stn. 2,5, 7, 8)
0.033
0.062
0.048
0.072
0.017
0.031
0.003
deviations.
Ecology 16 (1995) 213-222
Usually the patterns of enzyme activity showed a
ranking of aminopeptidase > esterase > chitobiase >
P-glucosidase > cy-glucosidase, and the mean ratios
of the highest potential activity rates of the enzymes
tested (usually found in the top horizon) relative to
o-glucosidase
were
the
activity
of
687/174/11/3/l
(Table 3). The amount carbon
potentially liberated by the enzymes in the O-l cm
sediment horizon was 421 f 58 mg C m-’ d- ’ for
the aminopeptidase,
89 + 65 mg for the esterase,
5.6 + 2.3 mg for the chitobiase, 1.7 f. 0.5 mg for the
P-glucosidase,
and 0.5 f. 0.03 mg for the a-glucosidase giving a total of 518 f 126 mg C mm2 d-‘.
Spatial or temporal variability (Table 6) was not
observed.
Parallel incubations
under a pressure of 1 atm
resulted in most cases in a depression of enzyme
activity of between 4-57% (Tables 4 and 51, but the
differences were mostly too small compared to the
data’s standard deviation so that barophilism (higher
activity under in situ pressure) could not be confirmed.
4. Discussion
To gain some insight into the spatial and temporal
variability of hydrolytic enzymatic activity in deep
sea sediment, stations in the West European Basin
of the NE-Atlantic
at I atm pressure
Esterase (stn. 1, 2, 3, 5, 7, 8)
4.909 f 2.165
2.865 f 0.798
2.511 f 0.859
2.428 k 0.556
2.347 + 1.517
1.811 + 1.222
0.765 + 0.453
P-Glucosidase (stn. 2, 5, 7)
cY-Glucosidase (stn. 2, 5, 7)
0.099 *
0.051 +
0.035 f
0.028 +
0.016 +
0.012 +
< 0.002
0.0296 f
0.0259 +
0.0211 +
0.0143 f
0.0064 f
0.0047 +
< 0.002
0.031
0.025
0.018
0.014
0.008
0.009
0.0029
0.0031
0.0028
0.0015
0.0007
0.0018
K. Poremba / FEMS Microbiology
Table 5
Hydrolytic
activities
h-
a (nmol cme3
Ecology 16 (1995) 213-222
‘1 in deep-sea sediments of the NE-Atlantic;
in situ pressure conditions
during incubation
Sediment depth km)
Aminopeptidase
o-1
l-2
2-3
3-4
4-6
6-8
8-10
Sediment depth km)
24.382 k 3.407
18.264 + 1.337
20.419 f 1.099
22.905 + 2.427
17.919 + 3.929
14.909 + 5.147
8.211 + 1.983
Chitobiase (stn. 2,5, 7,8)
7.707 + 5.603
4.076 + 1.892
3.026 + 1.026
2.324 + 0.666
2.167 f 0.816
1.723 + 0.959
0.754 + 0.311
P-Glucosidase (stn. 2,5, 7)
a-Glucosidase
o-1
l-2
2-3
3-4
4-6
6-8
8-10
0.325
0.166
0.102
0.078
0.039
0.040
0.041
0.098 f
0.130 f
0.077 f
0.044 +
0.016 f
0.013 +
< 0.002
0.0296 + 0.0029
0.0259 f 0.0031
0.0211 + 0.0028
0.0143 + 0.0015
0.0064 f 0.0007
0.0047 & 0.0018
< 0.002
a All values are means f standard
+
It
f
+
+
*
+
(stn. 2, 3, 5, 7, 8)
217
Esterase (stn. 1, 2, 3, 5, 7, 8)
0.134
0.091
0.063
0.058
0.019
0.041
0.039
Sediment depth km)
o-1
l-2
2-3
3-4
4-6
6-8
8-10
o-1
1-2
2-3
3-4
4-6
6-8
8-10
organic material. The heterogenity of POM settling
on the deep-sea floor as organic aggregates (marine
snow) is well known [23,24]. It is expected that a
significant part of proteinaceous matter in sediment
will be highly refractory [25]. The importance of the
specific composition of phytodetritus for its qualitity
as food for the deep-sea benthic community has also
recently been shown by a succession of utilization
of esterase activity @A) a and total bacterial
Spring: 46’N/17”W
TBN
11.43 * 1.21
5.22 f 0.46
3.23 10.19
2.94 + 0.24
3.34 f 0.34
3.59 * 0.35
1.18 f 0.18
Summer: 59”N/21”W
6.43 + 0.53
6.32 f 0.34
4.03 + 0.15
3.11 f 0.08
1.52 * 0.05
1.27 + 0.02
0.82 k 0.04
10.9
9.4
8.0
6.7
8.9
3.9
3.5
(stn. 5)
9.7
15.2
7.9
7.2
8.1
5.6
6.2
deviations.
number (TBN) in sediment of the NE-Atlantic;
Spring: 47”N/19”W
(sm. 1)
EA (nmol cmw3 h-r)
a All values are means f standard
(stn. 2,5,7)
deviations.
and in the Iceland Basin were visited in March/April
and July/August
1992.
The activity profiles mostly showed decreasing
gradients with increasing sediment depth, but the
slopes for individual
enzymes were different and
aminopeptidase
often had a small peak just below
the surface. This might indicate vertical shifts in both
the availability
and nutritive quality of degradable
Table 6
Temporal and spatial variation
conditions during incubation
0.034
0.106
0.062
0.038
0.008
0.007
(X
10’ cme3)
(stn. 4)
EA (nmol cme3 h- ‘)
4.76 + 2.66
1.29 f 0.25
1.32 + 0.12
1.79 + 0.28
1.29 f 0.14
0.88 f 0.26
0.29 k 0.03
Summer: 47”N/19”W
5.62 + 2.77
3.50 + 0.43
3.61 f 0.31
2.54 f 1.05
2.53 + 0.38
2.39 f 0.82
0.72 + 0.07
in situ pressure
TBN (X 10’ cmm3)
11.1
9.6
8.0
9.8
8.9
8.4
4.4
(stn. 7)
15.4
8.7
8.5
5.8
7.2
4.6
5.8
218
K. Poremba / FEMS Microbiology Ecology 16 (I 995) 213-222
rates during the microbial remineralization
of artifical or natural detritus by deep-sea sediment communities [26-281.
Alternatively,
the individual vertical gradients of
each enzyme may be due to a shift in the composition of organisms within the microbial community
that are actively producing enzymes. The enzyme
activities within the top 10 cm of the cores decreased
by 87-99%
of their surface values (only the
aminopeptidase
show a weak decline of 66%) while
the bacterial numbers decreased by 40-70%. This
led to individual profiles of per-cell activity for each
of the enzymes tested.
The patterns of hydrolytic enzymes showed a
dominance
of nitrogen-containing
polymers compared to N-free compounds indicated by the activity
ranking of aminopeptidase > esterase > chitobiase >
P-glucosidase > a-glucosidase
with ratios of about
687/174/11/3/l.
In contrast to this, a higher utilization activity of polysaccharides
relative to N-related polymers is typical in water samples (e.g.
Hoppe in the Kiel Bight [2]>. On the other hand,
particles collected in surface waters often show the
opposite. Smith et al. [9] sampled water and aggregates in coastal waters of the Southern California
Bight in 25 m water depth and measured aminopeptidase, chitobiase,
and a-glucosidase.
The authors
found ratios of about 300/10/l
in marine aggregates and about 4/1/l
in surrounding
sea water.
Recently Miiller-Niklas
et al. [29] studied the same
activities in coastal waters of the Adriatic Sea in
5-10 m water depth, and found ratios of 100/1/l
in
particles and 200/2/l
in ambient water. It is striking to note the dominance of N-related hydrolytic
activity over the polysaccharide degradation in water
samples relative to aggregates and the similarity
between activity ratios in sediment and in marine
snow. Small differences may be due to different
substrate concentrations
used (Miiller-Niklas
et al.:
2.5 pm01 I-‘, Smith et al.: 20 nmol I-‘, here:
0.05-l
pmol 1-l). Biogenic aggregates are known
as ‘hot spots’ of intense hydrolytic activity and are
the main vehicles for the downward flux of carbon
[15,30]. Aggregates possess enzymatic activities several orders of magnitude higher than surrounding sea
water, although their importance in the decomposition of organic material in the oceanic water column
is considered to be low [9,31,32]. However, the
general similarity of enzyme activity patterns in marine snow and sediment indicates the biochemical
relationship between POM of surface and deep water, and supports the hypothesis of pelagic-benthic
coupling in the oceans [1,33,34]. It seems that the
shift in the biochemical composition of organic material in offshore water columns, indicated by the
increasing C/N ratio with depth [35], is not contradictory to the faster decrease in polysaccharide mobilizing activity with depth compared to the activity
utilizing nitrogen containing macromolecules.
There are very few investigations
on hydrolytic
enzymes in deep marine sediments. Most included
fewer types of enzymes, and due to the lack of
standardized methods (substrate concentrations, incubation temperature, incubation duration, sample volume, or processing of the tested material during the
experiment), a direct comparision of activity rates is
difficult. A potential aminopeptidase
rate of lo-25
nmol cme3 hh ’ (top horizon) was found in marine
sediments from deep stations [lo] as well as from
shallow ones [21,22], indicating only a small impact
of environmental
conditions on the benthic readiness
for biological protein degradation. While the hydrolysis rates of chitobiase in surface waters are similar
to those of the P-glucosidase (around 5 pmol ml-’
hh’ [2]), their rates in sediment are different. In
shallow intertidal sediments /3-glucosidase activity
has found to be 4-times higher than the chitobiase
rate (1392 and 354 nmol cmp3 hh’ in the top
horizon, respectively)
[36]. In this study the same
ratio was shown between chitobiase and p-glucosidase, but about lOOO-times lower absolute rates.
The a-glucosidase
rate, which is usually higher in
surface waters than that of P-glucosidase
[2], was
the lowest activity of all enzymes tested in NEAtlantic sediment. This might reflect increased lability of starch under natural conditions
leading to
substantial depletion of the substrate for cr-glucosidases in POM deposited on the deep-sea floor.
Measurements
of bacterial
abundance
during
1983-1989 at one of our study sites (47”N 19”W)
have shown a doubling of benthic bacterial biomass
in July/August
compared to March [26]. Therefore,
seasonality was expected in our study, but neither the
measurements
of enzymatic activity nor those of
bacterial abundance showed seasonal variations (Table 6). Only the depth gradient of activity in spring
K. Poremba / FEMS Microbiology
was steeper than in summer, which suggests a small
seasonal influence on enzymatic activity. In addition,
slightly higher activities and cell numbers were found
at the sediment surface layer relative to deeper layers. The increases in biomass and activity were of
similar extent to those reported previously 1261, but
the variability of the values made detection of significant seasonality impossible. The lack of a seasonal
shift may be due to an unusually early planktonic
spring bloom in 1992 [37]. Moreover, it should be
noted that the assays reported here were at saturated
substrate concentrations
enabling the measurement
of a potential activity rate as opposed to the actual in
situ rate. It is possible that this method does not
resolve temporal variability, while assays performed
with natural substrate concentrations
would reflect
seasonal events. An assay of natural extracellular
enzyme activities was not possible, because the concentrations of specific biopolymers
at the deep-sea
floor or in sedimented POM are not known.
Spatial variability may be inferred, as increased
activities occurred at stations 1 and 5 (Table 6).
Station 1 lies below the carbonate compensation
depth (CCD), where the chemical solubilization
of
sedimented plankton shells might supply extraordinarily unprotected material at the top horizon. The
slightly elevated esterase rates between l-3 cm depth
in the sediments of stn. 5 might be due to a higher
input of POM at this site, since the water depth is
much shallower than the other stations. The difficulty of obtaining replicate sediment samples from
the various deep-sea stations has prevented statistically significant spatial differences from being detected in the same way as with seasonal variability.
Incubations performed under in situ pressure conditions were not significantly
different from those
performed at 1 atm. One should consider that hydrolytic activity is mostly extracellular,
and therefore
not necessarily closely correlated to the physiology
of the organisms producing the enzymes 1121. The
effect of pressure on structure and folding of isolated
peptide chains is small and even less compared to
temperature,
salinity or pH and their function as
biocatalysator is only slightly affected [38]. The typical abnormalities
of growth under unusual pressure
conditions like, e.g., elongated cell forms or reduction of protein biosynthesis
should be a results of
several synergistic effects within the cell [39]. There-
Ecology 16 (1995) 213-222
219
fore, extracellular hydrolytic activity might be a poor
indicator for barophilism, which usually is indicated
by a higher activity of deep-sea organisms under
elevated pressure [14].
The sum of carbon potentially liberated by the
different enzymatic
activities was compared with
sediment trap data from the area investigated. In the
top sediment horizon, 518 mg C mm2 dd’ would be
mobilized, which is about 200-times more than the
input of 2.74 mg particulate organic C me2 dd’ [33].
This discrepancy can be due to the use of saturated
substrate concentrations,
which may lead to an overestimation of the actual degradation rate. However,
the difference between the input of material and its
biological utilization may also support the idea of an
‘uncoupled hydrolysis’ as a biological strategy for
the benthic community in the deep sea. Recent investigations have shown the ability of the deep-sea
benthos to react rapidly to pulses of organic matter
[34,40-421, but not necessarily in a closely coupled
way. From that, it can be assumed that the detected
elevated potential activity of extracellular enzymes
indicate the readiness of the benthic microbial population to utilize incoming
organic material. The
proposition that extracellular
enzyme activity is a
key step in aquatic ecosystems [2-4,211 may be
particularly germane to the deep-sea benthic environment.
Acknowledgements
Thanks are due to the Meteor crews of the cruises
21/l and 21/6 for their excellent cooperation, Karen
Jeskulke for her technical assistance and Richard
Lampitt for discussing the manuscript.
This work
was supported by grant MUF 03F0565A of the
Bundesministerium
fiir Forschung
und Technik
(BMIW.
References
[r] Pfannkuche,
0. (1992) Organic carbon flux through the
benthic community in the temperate abyssal Northeast Atlantic. In: Deep-sea food chains and relation to the global
carbon cycle (Rowe, G.T. and Pariente, V., Eds.), pp. 183198. Kluwer Academic Publishers, Dordrecht.
220
K. Poremba/FEMS
Microbiology
121 Hoppe, H.-G. (1983) Significance of exoenzymatic activities
in the ecology of brackish water: measurements by means of
methylumbelliferyl-substrates.
Mar. Ecol. Prog. Ser. 11,
299-308.
[3] Meyer-Reil, L.-A. (1990) Microorganisms
in marine sediments: considerations
concerning
activity measurements.
Arch. Hydrobiol. Beih. 34, 1-6.
[4] Hoppe, H.-G. (1991) Microbial extracellular enzyme activity:
a new key parameter in aquatic ecology. In: Microbial enzymes in aquatic environments (Chrost, J., Ed.), pp. 60-79.
Springer-Verlag,
New York.
[5] Schniirer, J. and Rosswall, T. (1982) Fluorescein diacetate
hydrolysis as a measure of total microbial activity in soil and
litter. Appl. Environ. Microbial. 43, 1256-1261.
161 Hoppe, H.-G., Ducklow, H. and Karrasch, B. (1993) Evidence for dependency
of bacterial growth on enzymatic
hydrolysis of particulate organic matter in the mesopelagic
ocean. Mar. Ecol. Prog. Ser. 93, 277-283.
[7] Hoppe, H.G., Kim, S.J. and Gocke, K. (1988) Microbial
decomposition in aquatic environments: combined process of
extracellular enzyme activity and substrate uptake. Mar. Ecol.
Prog. Ser. 54, 784-790.
[8] Cho, B.C. and Azam, F. (1988) Major role of bacteria in
biogeochemical
fluxes in the ocean’s interior. Nature 332,
441-443.
[9] Smith, D.C., Simon, M., Alldredge,
A.L. and Azam, F.
(1992) Intense hydrolytic enzyme activity on marine aggregates and implications for rapid particle dissolution. Nature
359, 139-142.
[lo] Boetius, A. (1994) Microbial hydrolytic enzyme activities in
deep-sea sediments. HelgolEnder Meeresunters (in press).
[ll] Boetius, A. and Lochte, K. (1994) Regulation of microbial
enzymatic degradation of organic matter in deep-sea sediments. Mar. Ecol. Prog. Ser. 104, 299-307.
1121 Meyer-Reil, L.-A. and Kiister, M. (1992) Microbial life in
pelagic sediments: the impact of environmental
parameters
on enzymatic degradation of organic material. Mar. Ecol.
Prog. Ser. 81, 65-72.
[13] ZoBell, C.E. (1946) Marine microbiology,
240 pp. Cronica
Botanica Company, Waltham, MA, USA.
[14] Deming, J.W. (1986) Ecological strategies of barophilic bacteria in the deep ocean. Microbial. Sci. 3, 205-211.
[15] Lochte, K. and Turley, C.M. (1988) Bacteria and cyanobacteria associated with phytodetritus in the deep-sea. Nature 33,
67-69.
[16] Poremba, K. (1994) Simulated degradation of phytodetritus
in deep-sea sediments of the NE Atlantic (47”N, 19”W). Mar.
Ecol. Prog. Ser. 105, 291-299.
[17] Bianchi, A. and Garcin, J. (1993) Stratified waters the
metabolic rate of deep-sea bacteria decreases with decompression. Deep-Sea Res. 40, 1703-1710.
[18] Bianchi, A. and Garcin, J. (1994) Bacterial response to
hydrostatic pressure in seawater samples collected in mixedwater and stratified-water
conditions. Mar. Ecol. Prog. Ser.
111, 137-141.
[19] Barnett, P.R.O., Watson, J. and Conelly, C. (1984) A multiple corer for taking virtually undisturbed samples from shelf,
bathyl and abyssal sediments. Oceanol. Acta 7, 399-408.
Ecology 16 (1995) 213-222
[20] Hobbie, J.E., Daley, J.R. and Jaspar, S. (1977) Use of
Nuclepore filters for counting bacteria by fluorescence microscopy. Appl. Environ. Microbial. 33, 1225-1228.
[21] Meyer-Reil, L.-A. (1986) Measurement of hydrolytic activity
and incorporation of dissolved organic substrates by microorganisms in marine sediments. Mar. Ecol. Prog. Ser. 31,
201-206.
[22] Meyer-Reil, L.-A. (1987) Seasonal and spatial distribution of
extracellular enzymatic activities and microbial incorporation
of dissolved organic substrates in marine sediments. Appl.
Environ. Microbial. 53, 1748-1755.
[23] Alldredge, A.L. and Youngbluth,
M.J. (1985) The significance of macroscopic aggregates (marine snow) as sit& for
heterotrophic bacterial production in the mesopelagic zone of
the subtropical Atlantic. Deep-Sea Res. 32, 1445-1456.
[24] Alldredge, A.L., Cole, J.J. and Caron, D.A. (1986) Production of heterotrophic bacteria inhabiting macroscopic organic
aggregates
(marine snow) from surface waters. Limnol.
Oceanogr. 31, 68-78.
[25] Rice, D.L. (1982) The detritus nitrogen problem: new observations and perspectives from organic geochemistry.
Mar.
Ecol. Prog. Ser. 9, 153-162.
[26] Lochte, K. (1992) Bacterial standing stock and consumption
of organic carbon in the benthic boundary layer of the
abyssal North Atlantic. In: Deep-sea food chains and the
global carbon cycle (Rowe, G.T. and Pariente, V., Eds.), pp.
l-10. KIuwer Academic Publishers, Dordrecht.
[27] Pfannkuche, O., Lochte, K. and Thiel, H. (1988) Sedimentation of spring phytodetritus to the deep-sea floor. EOS 69,
1117.
[28] Turley. CM. and Loch@ K. (1990) Microbial response to
the input of fresh detritus to the deep-sea bed. Palaeogeogr.,
Palaeoclimatol.,
Palaeoecol. (Global and Planetary Change
Section) 89, 3-23.
[29] Miiller-Niklas, G., Schuster, S., Kaltenbiick, E. and Hemdl,
G.J. (1994): Organic content and bacterial metabolism in
amorphous aggregations of the northern Adriatic Sea. Limnol. Oceanogr. 39, 58-68.
[30] Honjo, S. (1982) Seasonality and interaction of biogenic and
lithogenic particulate flux at the Panama Basin. Science 218,
883-884.
1311 Rosso, A.L. and Azam, F. (1987) Proteolytic activity in
coastal ocean waters: depth distribution and relationship to
bacterial populations. Mar. Ecol. Prog. Ser. 41, 231-240.
[32] Karl, D.M., Knauer, G.A. and Martin, J.M. (1988) Downward flux of particulate
organic matter in the ocean: a
particle decomposition paradox. Nature 332, 438-441.
[33] Honjo, S. and Maganini, S.J. (1993) Annual biogenic particle
fluxes to the interior of the North Atlantic Ocean; studied at
34”N 21”w and 48”N 21”W. Deep-Sea Res. 40, 587-607.
[34] Graf, G. (1989) Benthic coupling in a deep-sea community.
Nature 341, 437-439.
[35] Martin, J.H., Knauer, G.A., Karl, D.M. and Broenkow, W.W.
(1987) VERTEX: Carbon cycling in the northeast Pacific.
Deep-Sea Res. 34, 267-285.
[36] King, G.M. (1986) Characterization of /3-glucosidase activity
in intertidal marine sediments. Appl. Environ. Microbial. 51,
373-380.
K. Poremba/FEMS
Microbiology
[37] Pfannkuche, O., Duinker, J.C., Graf, G., Henrich, R., Thiel,
H. and Zeitschel, B. (1993) METEOR Berichte, pp. 281,
Universitlt Hamburg 93-4.
[38] jaenike, R. (1988) Cellular components under extremes of
pressure and temperature.
In: Current perspectives in high
pressure biology (Jannasch, H.W., Marquis, R.E. and Zimmermann, A.M., Eds.), pp. 257-272. Academic Press, London, Toronto.
[39] Jaenike, R. (1988) Molekulare Mechanismen
der Adaption
von Bakterien an extreme Bedingungen.
Forum Mikrobiologie 10, 43.5-440.
Ecology 16 (1995) 213-222
221
[40] Gooday, A.J. (1988) A benthic foraminiferal response to the
deposition of phytodetritus
in the deep-sea. Nature 332,
70-73.
[41] Gooday, A.J., and Turley, C.M. (1990) Response by benthic
organisms to input of organic material to the ocean floor.
Philos. Trans. R. S. Lond. A. Math. Phys. Sci. 331, 119-138.
[42] Pfannkuche, 0. (1993) Benthic response to the sedimentation
of particulate
organic matter at the BIOTRANS
station,
47”N, 2O”W. Deep-Sea Res. 40, 135-1.50.