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Applied and Environmental Microbiology, September 1998, p. 3238-3245, Vol. 64, No. 9
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Nucleic Acid (DNA, RNA) Quantification and RNA/DNA Ratio
Determination in Marine Sediments: Comparison of Spectrophotometric,
Fluorometric, and HighPerformance Liquid Chromatography
Methods and Estimation of Detrital DNA
A.
Dell'Anno,1
M.
Fabiano,2
G. C. A.
Duineveld,3
A.
Kok,3 and
R.
Danovaro1,*
Faculty of Science, Marine Science,
University of Ancona, Ancona,1 and
Institute of Marine Environmental Sciences, University of
Genoa, Genoa,2 Italy, and
Netherlands
Institute for Sea Research, Texel, The Netherlands3
Received 10 November 1997/Accepted 4 April 1998
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ABSTRACT |
In this study, we compared three methods for extraction and
quantification of RNA and DNA from marine sediments: (i) a
spectrophotometric method using the diphenylamine assay; (ii) a
fluorometric method utilizing selective fluorochromes (thiazole orange
for total nucleic acids and Hoechst 33258 for DNA); and (iii) a
high-pressure liquid chromatography (HPLC) method which uses a specific
column to separate RNA and DNA and UV absorption of the nucleic acids
for quantification. Sediment samples were collected in the oligotrophic
Cretan Sea (eastern Mediterranean, from 40 to 1,540 m in depth) and
compared to the distribution and composition of the benthic microbial
assemblages (i.e., bacteria and microprotozoa). DNA concentrations
measured spectrophotometrically and by HPLC were not significantly
different, while fluorometric yields were significantly lower. Such
differences appear mainly due to fact that the stain-DNA complex is
strongly dependent on the DNA composition and structure. RNA
concentrations determined by the three methods displayed some
differences; fluorometric and spectrophotometric methods obtain RNA
concentration by difference and therefore may be biased by DNA
estimates. By contrast, the HPLC method provides independent
assessments of RNA and DNA concentrations. We tentatively estimated the
contribution of the detrital DNA to the total DNA pools in two ways.
The two calculations provided quite similar results indicating that the
majority of the DNA pool in the deep-sea sediments was detrital.
Microbial RNA generally accounted for almost the entire sedimentary RNA
pools below 100-m depth. RNA concentrations were found to decrease
along the Cretan shelf and slope. The RNA/DNA ratio calculated by using
fluorometric DNA concentrations was significantly correlated with
values of sediment community oxygen consumption only below 100-m depth
(dominated by the microbial biomass). These data suggest that the
RNA/DNA ratio, based on fluorometric estimates of DNA, can be used as an indicator of benthic metabolic activity, but only when metazoan contribution to the microbial DNA is negligible.
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INTRODUCTION |
Nucleic acid determination in marine
habitats has attracted special attention because of the relationship
between RNA/DNA ratios and the growth rates of a wide variety of marine
organisms such as phytoplankton (3, 4, 12, 13, 44), bacteria (22, 24, 33), invertebrates (42), and fish
(5, 6). Other studies of natural seawater and sediment
samples have been undertaken to investigate the relationships between
RNA/DNA ratio and the metabolic state of the microbial communities
associated with suspended and sedimentary organic particles (9,
16). However, Jeffrey et al. (24) found no correlation
between RNA/DNA ratios of natural bacterioplankton communities and
other measures of metabolic activity or growth (thymidine and leucine
incorporation). There are several problems which may compromise the
interpretation of RNA/DNA ratios in intact sediments as a measure of
benthic community metabolism. The most important of them is the
presence of substantial amounts of DNA associated with dead cells
and/or absorbed to particles (i.e., detrital DNA) in the DNA pools in the sediments and in the water column as well. Holm-Hansen et al.
(21) first tried to use DNA concentrations as a measure for
living biomass in the oceans; they found unreasonably high concentrations and postulated that most of the DNA was not associated with living cells and thus not immediately degraded. Also, Danovaro et
al. (9) found a large fraction of the DNA in sediments from the deep Mediterranean to be unaccounted-for bacterial standing stocks.
The significance and nature of this apparently detrital DNA is still
uncertain (1).
For the determination of nucleic acids from sediments, two methods are
generally used: spectrophotometric, based on the specific absorbance of
nucleic acids (9, 18), and fluorometric, using specific
fluorescent stains (32, 45). To date, comparative studies
for nucleic acid determination have been performed with bacteria, algal
cultures, and natural seawater samples (25, 37), but there
are no examples of similar comparisons for nucleic acid determination
in marine sediments.
In this study, for measuring RNA and DNA in marine sediments, we
applied three methods which differ with respect to the nucleic acid
extraction procedure. Moreover, two of the methods, referred to here
for simplicity as the spectrophotometric and high-pressure liquid
chromatography (HPLC) methods, use spectrophotometric quantification of
nucleic acids, while the third (the fluorometric method) uses fluorescent dyes. The three procedures share the following advantages: (i) the extraction time is relatively short, allowing the processing of
the large number of samples usually collected in field studies; and
(ii) quantification of DNA and RNA in the same subsamples is possible,
thus allowing for determination of the RNA/DNA ratio. The aims of this
study were to (i) compare the results of nucleic acid determination
using the above-named three methods; (ii) estimate the detrital
fraction of the DNA and RNA pools in the Cretan sediments by
calculating the DNA and RNA contribution by intact and stainable microbes (i.e., bacterial and protozoan densities by microscopy); and
(iii) explore the relationships between the patterns of RNA/DNA ratios
and sediment community oxygen consumption (SCOC).
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MATERIALS AND METHODS |
Study area and sampling.
Sediment sampling was carried out
on the continental shelf and slope of the north coast of Crete and in
the adjacent deep basin of the Cretan Sea (south Aegean, eastern
Mediterranean). This area is one of the world's most oligotrophic
seas, with a primary production as low as 20 to 30 g of C
m
2 year1 (14). The Cretan Sea is
further characterized by high bottom-water temperatures (13 to
14.5°C) and strong water column stratification.
Undisturbed sediment samples were collected by using a multicorer in
September 1995 at seven stations situated along a transect of depths
from 40 to 1,540 m: 40, 100, 200, 500, 700, 900, and 1,540 m (Fig.
1). For nucleic acid determination, we
used the top 1-cm slices of two cores, which were subsequently
homogenized and deep frozen at 
20°C. For bacterial and protozoan
counting, three to five replicate subsamples (0.63 cm3)
were collected with sterile cutoff syringes from the same cores used
for nucleic acid analysis. The subsamples were fixed with filtered
(0.2-µm-pore-size filter) seawater containing 2% buffered formalin
and stored at 4°C for later analyses in the laboratory.
Nucleic acid analysis.
Before analysis, larger macroscopic
organisms were removed from the samples. All of the materials used for
nucleic acid analysis were carefully cleaned by soaking in 1 N
NaOH-10% HCl-MilliQ water to remove organic matter contamination and
subsequently treated as described by Moran et al. (32) to
avoid nuclease contamination. All of the solutions were prepared with
MilliQ water and then autoclaved. Amounts of DNA and RNA were
determined by the spectrophotometric, fluorometric, and HPLC methods.
For each method, internal standards of calf thymus DNA and baker's
yeast RNA (5 to 10 µg) were added to replicate subsamples before
extraction. The final yields of the internal standards of DNA and
RNA were on average 60 and 85% for the spectrophotometric method, 55 and 80% for the fluorometric method, and 95 and 90% for the HPLC
method. DNA and RNA concentrations in the sediments were not corrected
for percent recovery of the internal standards and were calculated from
calibration curves of calf thymus DNA and baker's yeast RNA prepared
according to each analytical protocol. Data were normalized to sediment
dry weight after desiccation (60°C, constant weight).
(i) Spectrophotometric method.
Nucleic acid extraction and
measurement were done by the procedures of Zachleder (50) as
applied by Danovaro et al. (9) and Danovaro (10),
with a few modifications to enhance DNA extraction from the sediment.
Briefly, 1 g of sediment (three replicates) was treated with 3.0 ml of 0.5 N perchloric acid, stirred for 3 min, and sonicated three
times for 1 min (with intervals of 30 s). Nucleic acid extraction
was carried out at 75°C for 30 min under continuous stirring. After
centrifugation (3,000 × g, 10 min), the absorbance of
the total nucleic acid content (TNA) in the supernatant was measured at
260 nm. DNA absorbance was determined with a diphenylamine (2% in
acetic acid) light-activated reaction (40 W, 12 h) at 598 nm and
converted to concentration, using standard solutions of calf thymus
DNA. DNA concentration was then reported as equivalent of absorbance at
260 nm in order to calculate by difference the absorbance due to RNA:
ABSRNA = ABSTNA ABSDNA, where
ABSRNA is the absorbance of RNA, ABSTNA is the
absorbance of TNA, and ABSDNA is the absorbance of DNA. RNA
absorbance (260 nm) was then converted to concentration, using standard
solutions of baker's yeast RNA. Since TNA absorbance at 260 nm might
be affected by the interference due to inorganic compounds, we used
sediment subsamples, previously treated in a muffle furnace (550°C,
4 h) as blanks. Sensitivity of the method has been tested on DNA
and RNA standards (accuracy of ±1.0 µg) and appeared to be adequate
for field investigations.
(ii) Fluorometric method.
Initially, nucleic acid extraction
for the fluorometric determination was performed with sediments
homogenized in Tris-Ca2+ buffer. This buffer provided the
best recovery and the lowest variability in natural seawater samples
(3). However, we found that this procedure was not effective
in the extraction of both RNA and DNA from sediments. Therefore, to
extract nucleic acids, we used sodium dodecyl sulfate (SDS), which is
efficient in cellular lysis and is commonly used in protocols for
extracting nucleic acids from marine sediments (35, 45).
Nucleic acid extraction for fluorometric determination was carried out
as follows: 1 g of sediment (three replicates) was added to 2 ml
of 0.2 M sodium phosphate buffer containing 0.1 M EDTA (pH 8.0),
stirred for 5 min, and then mixed with 2 ml of SDS (10%). The samples
were incubated for 2 h at 65°C and subsequently subjected to
repeated freeze-thaw steps (five times, from 65 to 80°C). Nucleic
acids were extracted from the sediment by centrifuging the lysate at
12,000 × g at 10°C for 15 min. The supernatant was
transferred to another tube; the pellet was resuspended in 2 ml of
sodium phosphate buffer, incubated at 65°C for 15 min, and
centrifuged as described above. This supernatant was added to the first
and dialyzed against sterile water to remove low-molecular-weight
contaminants such as SDS and salts. The volumes of the final dialysate
were measured, added to absolute ethanol (ethanol volume = 3 × dialysate volume), and stored (80°C, 2 h). After
centrifugation (10,000 × g, 20 min) to precipitate
nucleic acids, the pellet was washed with ethanol 70%, centrifuged
again (10,000 × g for 20 min), dried under
N2, and resuspended in 100 µl of MilliQ water for
fluorometric analysis. Before fluorometric analysis, an aliquot of the
resuspension was analyzed by gel electrophoresis to ensure the presence
of nucleic acids in the extract (Fig. 2).
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FIG. 2.
Agarose gel electrophoresis of sedimentary nucleic acids
by extraction using the fluorometric method. Lanes: 1, lambda plasmid
digested with HindIII (molecular weight marker); 4 and
5, calf thymus DNA standard (0.4 and 0.8 µg, respectively); 7, sediment sample from 1,540-m depth; 8, sediment sample from 100-m depth
containing internal standard of calf thymus DNA, treated with RNase; 9, sediment sample from 100-m depth treated with RNase.
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Subsamples (25 µl) were then analyzed with two fluorescent dyes,
thiazole orange for TNA (
26) and Hoechst 33258 (Hoechst)
for
DNA (
37), using the procedure described by Berdalet and
Dortch (
3). Fluorescence was measured with a Perkin-Elmer
LS50B
spectrofluorometer (thiazole orange, 511-nm excitation and 533-nm
emission; Hoechst, 360-nm excitation and 460-nm emission). DNA
concentrations were calculated from calf thymus DNA standards
stained
with Hoechst. The same DNA standards were stained with
thiazole orange
to determine the DNA contribution to the total
thiazole orange
fluorescence. The RNA contribution in the thiazole
orange fluorescence
(FTO
RNA) was estimated as FTO
RNA = FTO
TNA FTO
DNA, where FTO
TNA is
the fluorescence of TNA after staining
with thiazole orange and
FTO
DNA is the calculated contribution
of DNA in the
thiazole orange fluorescence (as determined from
Hoechst staining). RNA
concentrations were calculated from baker's
yeast RNA standards after
thiazole orange staining.
In addition, an aliquot of the samples was treated with RNase and
stained with thiazole orange and Hoechst to compare the
fluorescence
yields obtained using the two stains on the same
DNA matrix (see
Discussion for comments on results).
(iii) HPLC method.
For HPLC determination of RNA and DNA,
0.2 g of sediment (three replicates) was added to 1 ml of Tris-HCl
buffer (with 2% SDS and 10% EDTA) and sonicated (Soniprep 150 MSE)
three times (10 s with 1-min interval). After sonication, the samples
were centrifuged for 5 min (3,000 × g), and the
supernatant was filtered over a 0.45-µm-pore-size cellulose acetate
filter to remove particulate material. All manipulations were carried
out at room temperature. The filtrate was directly injected into the
HPLC system, which consisted of a Waters 600E pump and controller unit,
connected via a Nucleogen 4000-7 DEAE anion-exchange column
(Machery-Nagel) (8) to a Waters 994 photodiode array. In
accordance with the method of Coppela et al. (8), we used a
urea buffer with a KCl gradient. The elution gradient and the flow rate
(Table 1) were adapted to overcome
problems caused by precipitation of the Tris-HCl extraction buffer
(with SDS and EDTA) when mixed with the urea buffer. Unretained
material (proteins) produced a large peak at the start of the
chromatogram. Figure 3 shows the
chromatogram of sedimentary DNA and RNA. The identity of the peaks in
the chromatogram was confirmed by inspection of the absorbance spectrum
(DNA and RNA maximum absorbance at 260 nm) in combination with either
coinjection of standards (calf thymus DNA and baker's yeast RNA) or
digestion of RNA with RNase. The areas of DNA and RNA peaks were
integrated at 260-nm wavelength. Peak areas were converted to
concentrations by using calibration curves obtained from standard
solutions of calf thymus DNA and baker's yeast RNA.
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TABLE 1.
Gradient flow rate and relative composition of eluents
used for HPLC determination of nucleic acids at a gradient flow rate of
1.6 ml/min
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Bacterial and protozoan analyses.
For bacterial analysis,
subsamples were sonicated three times (Sonifier Branson 2200, 60 W for
1 min), diluted 100 times, stained with acridine orange (0.01%, final
concentration) and filtered on black Nuclepore 0.2-µm-pore-size
filters. This procedure appeared to be the most appropriate both for
cell recovery and for normalization of data to sediment dry weight
(31). The filters were analyzed under epifluorescence
microscopy (Zeiss Universal microscope). Bacterial DNA and RNA were
estimated by assuming conversion factors of 3.3 fg of DNA
(41) and 4.2 fg of RNA (19) for cells of the same
size encountered in this study. Data were normalized to sediment dry
weight after desiccation (60°C, constant weight).
For protozoan (heterotrophic flagellates) analysis, the subsamples were
diluted in prefiltered (0.2-µm-pore-size filter) seawater
containing
acridine orange (0.01%, final concentration) to stain
the DNA and
filtered onto 2.0-µm-pore-size black-stained Nuclepore
filters. The
filters were subsequently stained as described by
Bak and Nieuwland
(
2) and scanned at a magnification of ×1,000.
Protozoan DNA
and RNA were estimated by assuming conversion factors
of 3.2 pg of DNA
cell
1 and 4.4 pg of RNA cell
1 (as average
of the values reported for different protozoan species
30). Data were normalized to sediment dry weight
after desiccation
(60°C, constant weight).
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RESULTS |
Spectrophotometric method.
DNA and RNA concentrations and
RNA/DNA ratios are reported in Table 2.
DNA concentrations ranged from 39.7 ± 14.2 at 40-m depth down to
12.3 ± 8.2 µg g1 at 1,540-m depth. The average
DNA concentration (± the standard deviation [SD]) was 30.9 ± 10.6 µg g1 (median, 37.4 µg g1). The
RNA concentration strongly decreased with water depth, from 29.9 ± 5.0 at 40 m to 0.4 ± 0.7 µg g1 at
1,540 m, with an average of 11.1 ± 10.9 µg
g1 (median, 5.1 µg g1). The RNA/DNA
ratios showed a similarly decreasing pattern with increasing water
depth and ranged from 0.96 to 0.004 (100 and 900 m, respectively).
Fluorometric method.
DNA concentrations ranged from 18.6 ± 0.3 (40 m) to 31.4 ± 1.9 µg g1 (100 m) in the
shelf (<200-m depth) and from 8.4 ± 0.7 (700 m) and 2.1 ± 0.5 µg g1 (1,540 m) in the slope and deep basin (Table
2). The average DNA concentration (±SD) was 10.3 ± 10.1 µg
g1 (median, 5.1 µg g1). Also, the highest
RNA concentrations (25 ± 3.5 µg g1) were found at
100-m depth, whereas in the other stations, RNA concentrations varied
in a narrow range (from 3.6 ± 0.3 to 7.5 ± 1.9 µg
g1 [Table 2]). The average RNA concentration was
8.4 ± 6.9 µg g1 (median, 5.7 µg
g1). The RNA/DNA ratio showed an increase with depth,
from 0.3 at 40-m depth to 2.15 at 1,540-m depth.
HPLC method.
DNA concentrations showed relatively little
variation, ranging between 39.8 ± 4.4 µg g1 at
40-m depth and 25.2 ± 4.3 µg g1 at 900-m depth
(Table 2). The average DNA concentration (±SD) was 33.6 ± 4.3 µg g1 (median, 33.6 µg g1). In contrast
to DNA, concentrations of RNA showed a clear decrease with depth, from
24.1 ± 2.4 µg g1 at 40-m depth to 1.3 ± 0.2 µg g1 at 1,540-m depth (Table 2). The average RNA
concentration was 7.0 ± 7.4 µg g1 (median, 2.9 µg g1). The RNA/DNA ratios showed a similar decreasing
trend, from 0.6 to 0.04 (depths of 40 and 1,540 m, respectively).
Bacteria and protozoa.
The density distribution of bacteria in
sediment samples is reported in Table 3.
The highest bacterial density (3.48 × 108
g1) was observed at 100-m depth. From 100 m
downward, bacterial densities showed a general decline with increasing
water depth, reaching the lowest value at 1,540 m (1.02 × 108 g1). The average bacterial density was
2.66 × 108 g1.
As for bacterial density, protozoan density (Table
3) was low at the
shallow 40-m station and increased toward the shelf
edge (200 m), where
highest densities (14.4 × 10
5 g
1) were
found. At greater depths, protozoan densities dropped again,
reaching a
level of 8.9 × 10
5 g
1 at 1,540 m. The
average protozoan density was 11.12 × 10
5
g
1.
The concentration of bacterial DNA estimated on the basis of conversion
factors (see Materials and Methods) ranged from 1.15
to 0.34 µg of
DNA g
1, and that of bacterial RNA ranged from 1.46 to
0.43 µg of RNA
g
1 (Table
3). The average concentrations
of bacterial DNA and RNA
were 0.88 and 1.12 µg g
1,
respectively. The estimated concentration of protozoan DNA varied
between 2.73 and 4.59 µg of DNA g
1, and that of
protozoan RNA ranged between 3.75 and 6.31 µg of
RNA
g
1 (Table
3). The average concentrations of protozoan DNA
and RNA
were 3.56 and 4.89 µg g
1, respectively.
Using the estimates of nucleic acid concentrations derived from
microscopic counts, we calculated DNA and RNA microbial (bacterial
plus
protozoan) contribution to the total pools of the two nucleic
acids
determined by the different methods (Table
3). Microbial
DNA of intact
cells (as determined by epifluorescence microscopy)
accounted for 16, 90, and 13% of the total DNA concentrations
as measured by the
spectrophotometric, fluorometric, and HPLC
methods, respectively.
Microbial RNA accounted on average for
332, 99, and 174% of the total
RNA concentrations determined spectrophotometrically,
fluorometrically,
and by HPLC, respectively. In many samples,
especially those from
deeper stations, the estimated microbial
RNA concentration greatly
exceeded the total RNA concentrations
measured spectrophotometrically
and by HPLC. The contribution
of microbial RNA to the RNA pools
determined fluorometrically
surpassed 100% only at depths of 200 and
500 m but was close to
90% in the other deep-sea stations.
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DISCUSSION |
Comparison of methods for DNA estimates.
DNA concentrations
measured spectrophotometrically and by HPLC in the seven stations of
the Cretan Sea were not significantly different (analysis of variance
[ANOVA] with replication; F = 0.89, P = 0.35 [i.e., all stations with all replicates]). By contrast, fluorometric DNA concentrations were significantly different, i.e.
lower, than spectrophotometric and HPLC measurements (ANOVA; F = 49.3, P < 0.01 and
F = 822.75, P < 0.01, respectively).
Possible explanations for these discrepancies concern the extraction
phase as well as the detection phase of the three methods. The
importance of the extraction step is illustrated by a comparison that
we made between the DNA yields obtained by the fluorometric method and
by the procedures of Tsai and Olson (45) and of Thiel and Higgins (43) (both used a fluorometric determination). The
results obtained with the Tsai and Olson (45) method,
specific for sedimentary DNA isolation, were very close to those of our
fluorometric method (Fig. 4a), while the
simple mortar homogenization in the Thiel and Higgins (43)
method yielded 10 to 20-times-lower DNA concentrations (data not shown)
even though it most likely causes the least damage to DNA molecules.
Leff et al. (28) compared different extraction procedures
(23, 34, 45) on stream sediments and recommended the Tsai
and Olson (45) procedure if purity and the presence of
eukaryotic DNA are of concern (as in our case) but found highest yields
with the method of Ogram et al. (34). The percentages of
internal DNA standard recovered by our spectrophotometric and fluorometric methods were almost equal, suggesting that there is little
difference between the extraction efficiencies of dissolved (i.e.,
standard) DNA. This, however, does not imply that the methods are also
equally efficient in the release of bound DNA (in cells or particles).
However, because we found similar DNA values with the Tsai and Olson
(45) method and our fluorometric method, there is little
reason to assume that our method is not efficient in the release of
DNA. Therefore, extraction alone cannot explain the lower fluorometric
yields.
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FIG. 4.
Comparison of two methods for fluorometric determination
of DNA. (a) Comparison with the Tsai and Olson (45)
extraction procedure; (b) comparison between DNA measured with thiazole
orange after treatment with RNase (HTO-DNA) and with Hoechst without
RNase treatment (HO-DNA).
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A more likely explanation for the significant DNA yield discrepancies
is that while the diphenylamine and HPLC determinations
are relatively
unaffected by DNA base composition and DNA molecular
structure
(
25), fluorescence yield of the Hoechst-DNA complex
is
strongly dependent on the composition and structure of DNA.
Hoechst
fluorescence is higher in AT-rich DNA than in GC-rich
DNA
(
33). Therefore, when DNA is quantified with Hoechst,
concentrations
appear to be affected by differences between the base
composition
of the sample DNA and the standard DNA (calf thymus; 60%
A + T
[
7]). In this regard, Paul and Myers
(
37) assumed for natural
heterogeneous bacterial communities
the same percent A + T of
the calf thymus DNA. However, if the
actual microbial A + T content
in our sediments were lower, this
method would underestimate the
concentration of microbial DNA.
Furthermore, Hoechst is a groove-binding DNA ligand that becomes
brightly fluorescent when it binds to the double-stranded
B form of the
DNA (
29) that represents the biologically active
form of
DNA. Paul (
36) reported that denatured DNA gives a low
contribution to the Hoechst-DNA fluorescence, while Fara et al.
(
17) found that DNA from seawater samples, after treatment
with
DNase, produced a negligible fluorescence with Hoechst and an
80%
reduced fluorescence with thiazole orange. We measured the
Hoechst
fluorescence yields with sedimentary DNA extracted in
perchloric acid
(as in the spectrophotometric protocol in which
the diphenylamine
reacts with hydrolyzed DNA) and adjusted to
pH 7.8, and we found a
fluorescence similar to the background
signal, meaning that Hoechst did
not bind the hydrolyzed DNA.
It is therefore likely that when the DNA
molecule structure is
altered, formation of the Hoechst-DNA complex is
compromised and
thus the fluorescence is greatly reduced.
The different affinities of Hoechst and diphenylamine with respect to
hydrolyzed DNA could explain differences found in our
samples and in
other natural seawater samples assessed fluorometrically
and
spectrophotometrically (
25,
37,
38). Therefore, it might
be
expected that in natural samples containing large amounts of
denatured,
damaged, and/or hydrolyzed DNA (i.e., detrital DNA),
Hoechst
measurements give yields lower than those obtained by
the diphenylamine
assay or by HPLC.
Comparison of methods for RNA estimates.
Estimates of total
RNA in natural samples are difficult because of the lability of the RNA
molecule (35). Since RNA concentrations determined by
fluorometric and spectrophotometric methods are calculated by
difference, their RNA estimates could be biased by the TNA and DNA
estimates. By contrast, the HPLC method, because of independent
assessments of RNA and DNA, is not biased by the DNA determination. RNA
concentrations obtained with spectrophotometric and HPLC methods were
not significantly different at most stations (ANOVA; F = 2.28, P = 0.14); only at depths of 100 and 200 m
were spectrophotometric values significantly higher than HPLC values (ANOVA; F = 5.17, P < 0.05). By
contrast, RNA concentrations determined fluorometrically were higher
than those obtained by HPLC (ANOVA; F = 6.878, P < 0.05) but not significantly different from those obtained by the spectrophotometric method (ANOVA; F = 3.284, P = 0.08). As for DNA, we tested the
fluorometric method for the recovery of RNA internal standards. This
resulted in a mean recovery of 80% of the added RNA, indicating that
no major loss or destruction of RNA occurred during the extraction
procedure. Therefore, no special protocols for RNA stability (i.e., use
of guanidium isothiocyanate and diethyl pyrocarbonate treatment of
glassware and solutions) appear to be necessary. However, a
complication may arise in the RNA determination with the fluorometric
method because of the different affinities of Hoechst and thiazole
orange for altered or damaged DNA. Figure 4b shows the DNA
concentrations in extracts of the same sample, one stained with Hoechst
and the other stained with thiazole orange (the latter previously
treated with RNase). Staining with Hoechst clearly gave much lower DNA
yields. As a consequence, RNA concentrations would be overestimated in
this case. As we do not know the contribution of the damaged DNA to the
fluorescence detected with thiazole orange, thiazole orange fluorescence cannot be used for correction of the DNA and RNA estimates.
Finally, RNA concentrations measured fluorometrically may be
particularly affected by the kind of standard used for calibration.
In
this regard, Mordy and Carlson (
33) reported that especially
when low RNA concentrations are present, different RNA standards
produce different fluorometric yields: the use of mRNA standards
may
produce overestimates compared to the use of rRNA or underestimates
compared to the use of tRNA. However, since changes in the relative
significance of ribosomal, transfer, and messenger RNAs may occur
in
marine sediments, as a function of the metabolic state of the
living
cells, these indications must be further investigated.
Comparison of methods for assessing the detrital fraction of DNA.
Large amounts of DNA are supplied to the benthos through
sedimentation of particles produced in the photic layer (1,
48). This "rain" of particle-associated DNA can serve as a
potentially important source of labile organic N and P and/or may be
used as a reservoir of nucleic acid precursors that are energetically expensive to synthesize ex novo (39). Especially in deep-sea benthic habitats, characterized by low inputs of labile organic matter,
detrital DNA could represent a suitable and high-quality food source
for heterotrophic metabolism. To evaluate the ecological role of the
detrital DNA, quantitative estimates of this fraction are needed.
However, the actual amounts of detrital DNA are not easy to assess.
Winn and Karl (49) at various Pacific Ocean locations and
Bailiff and Karl (1) in the Antarctic Peninsula region
provided the first estimates for the relative significance of
particulate detrital DNA to the total DNA (75 to 90 and 0 to 93%,
respectively). These estimates were obtained indirectly by converting
both total DNA (as detrital plus living organic matter) and ATP (living
organic matter) concentrations into carbon equivalents and subsequently
subtracting these quantities to obtain estimates of detrital DNA. Using
the same approach and conversion factors, we calculated the
contribution of the living fraction to the total DNA pool (ATP data
were summarized from reference 46), but we found
extremely low values (on average 1.6%, ranging from 0.18 and 3.3% at
depths of 100 and 900 m, respectively). The estimated amount of
living carbon depends on the conversion factor C/ATP (here 250, used
also by Bailiff and Karl [1]), but Graf and Linke
(20) reported that C/ATP ratios in deep-sea sediments may
range from <200 to >2,000. Therefore, the use of the C/ATP conversion
factor appears too variable to provide accurate estimates.
An alternative approach is to estimate the detrital DNA by difference
between total DNA and the contribution of the living
components (i.e.,
microscopically visible intact and stainable
bacteria and protozoa). We
calculated microbial DNA contribution
to the total DNA pool for the
following reasons: (i) bacteria
and protozoa account for more 90% of
the total biomass in deep-sea
samples of the Cretan Sea (
11,
47), and (ii) macroscopic metazoans
were removed from our samples
before analysis. We found average
microbial DNA contributions to the
total DNA pool of 16 and 13%,
respectively, for spectrophotometric and
HPLC total DNA determinations
(Table
3). Hence, with these methods, 84 and 87% of the DNA extracted
from the sediment was either from
organisms that were not counted
microscopically or from the detrital
pool.
By contrast, with fluorometric determinations, bacterial and protozoan
DNA from intact cells below 100-m depth generally accounted
for the
whole DNA pool (Table
3), whereas at shallow depths (i.e.,
40 and
100 m) they accounted on average for only 17.5%. These
results
demonstrate that when benthic biomass is dominated by
microbes (i.e.,
bacteria and protozoa together accounting for
>90% of total biomass,
such as below 100-m depth [
11]), Hoechst-DNA
concentrations are very close to those estimated from the microscopic
counts of intact cells. Conversely, at shallow depths, other DNA
pools
contributed to the fluorometric DNA estimates.
Estimates of detrital DNA calculated by difference between total and
microbial DNA pools must be viewed with caution for the
following
reasons: (i) the estimates may be biased by the use
of conversion
factors (though to a much lower extent than for
ATP); (ii) since no
conversion factors are actually available
for benthic communities, we
used factors obtained from planktonic
bacteria and protozoa; (iii) the
conversion factors depend on
cell biomass, whereas we estimated DNA
content from cell number
(though from similar cell sizes); and (iv)
microbial assemblages
display seasonal changes in DNA content
(
27).
Finally, to provide additional support for these estimates of detrital
DNA, we calculated the percentage contribution of the
fluorometric DNA
concentrations (assumed to represent DNA of intact
cells) to the DNA
pool measured spectrophotometrically and by
HPLC (assumed to represent
the microbial-plus-detrital DNA fraction).
DNA fluorometric yields
accounted on average for about 30% of
the spectrophotometric or HPLC
DNA concentrations (Table
2).
The resulting 70% of DNA concentration
not detected fluorometrically
was roughly close to 84 to 87% of the
total spectrophotometric
and HPLC DNA pool unaccounted by microbes. The
major differences
between these estimates were observed in the
continental shelf
at shallow depths (<100 m) where the fluorometric
DNA concentrations
gave highest yields, accounting on average for about
65% of spectrophotometric
or HPLC DNA concentrations. Such a high
contribution seems to
confirm that at depths of 40 and 100 m,
relatively large amounts
of nonmicrobial DNA are present (as suggested
by the much higher
meiofauna and macrofauna biomass [
11,
47]).
Further investigations are needed to confirm these data, but so far our
results indicate that the combined use of a spectrophotometric
or HPLC
and Hoechst fluorometric DNA assessment could represent
an alternative
approach to estimate the detrital DNA fraction
in the sediments.
RNA/DNA ratio in relation to environmental constraints.
The
decrease of RNA concentrations along the Cretan shelf and slope
reflects a drop in the benthic metabolic activity; Duineveld et al.
(15), from synoptic measurements on SCOC, demonstrated a
clear decrease of the respiratory activity with increasing depth. Since
below 100-m depth meiofauna and macrofauna accounted together for less
than 10% of the total benthic biomass (11, 47), most of the
respiratory activity in deeper sediments will be microbial (15,
40). The dominance of microbes is consistent with the very high
proportion of microbial RNA, which generally accounted for almost the
entire sedimentary RNA pools below 100-m depth (Table 3). This finding
implies that changes in the microbial metabolic activity in the
sediment should be reflected by sedimentary RNA concentrations measured
by chemical methods.
The RNA/DNA ratio in bacteria and phytoplankton has long been thought
to be proportional to levels of metabolic activities
and growth rates
(
12,
13). In a study on natural water samples,
Jeffrey et
al. (
24) found that RNA and DNA were strongly correlated
with bacterial numbers but also that the RNA/DNA ratio did not
correlate with either bacterial growth or metabolic activity and
concluded that this ratio cannot be used as biochemical indicator
of
activity in the environment. Correlation between RNA/DNA ratio
and
bacterial metabolic activity in natural samples could be biased
by the
presence of detrital DNA because its proportion is not
constant in time
and space. To test the reliability of the use
of the RNA/DNA ratio as
an indicator of the benthic metabolic
activity, we tested the
correlation between SCOC and the ratios
of RNA to DNA, calculated
between RNA determined spectrophotometrically
or by HPLC (as
fluorometric values might be overestimated as previously
discussed) and
DNA determined fluorometrically (which is assumed
to represent the
biologically active fraction of the total DNA
pool). Figure
5 illustrates the significant correlation
found
between the two RNA/DNA ratios and SCOC below 100-m depth
(
P <
0.01). When the shallowest stations (i.e., 40- and 100-m depth)
are included, the correlation between SCOC and RNA/DNA
ratios
becomes weak. This appears to be due to the higher meiofauna and
macrofauna oxygen consumption at shallow stations (
11,
47).
Interestingly, a highly significant correlation between sedimentary
RNA
content (measured spectrophotometrically and by HPLC) and
SCOC was
found (
r = 0.875,
P < 0.01 [spectrophotometric] and
r = 0.997,
P < 0.01 [HPLC]), indicating that RNA concentrations
alone might
predict better than the RNA/DNA ratio the benthic
metabolic activity.
View larger version (20K):
[in this window]
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|
FIG. 5.
Relationships between SCOC and RNA/DNA ratios (based on
spectrophotometric or HPLC determinations of RNA and fluorometric
determinations of DNA) in the deep-sea sediments (>100-m depth) of the
Cretan Sea.
|
|
These data suggest that the RNA/DNA ratio, calculated as described
above, could be used as an indicator of the metabolic activity
of
heterogeneous benthic microbial assemblages. However, the use
of this
ratio as indicator of benthic metabolism appears to be
hampered when
other (nonmicrobial) contributions to the DNA are
not negligible.
| |
ACKNOWLEDGMENTS |
We thank the crew and officers of the R.V. Philia for
help during sampling. We thank A. Tselepides and A. Eleftheriou
(Institute of Marine Biology of Crete) for great collaboration and
providing laboratory facilities. We thank also Rita Colwell (Baltimore) and Elisa Berdalet (Barcelona) and two anonymous referees for suggestions on improving an early draft of the report.
This research was undertaken in the framework of the CINCS project. We
acknowledge support from the European Commission's Marine Science and
Technology Program (MAST II) under contract MAS2-CT-940092.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Marine Science,
Facoltà di Scienze, Università di Ancona, Via Brecce
Bianche, 60131 Ancona, Italy. Phone: 39 71 2204652. Fax: 39 71 2204513. E-mail: danovaro{at}popcsi.unian.it.
| |
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Abstract
Introduction
