Summary of currently in use primary production filtration and acidification methods. POC = particulate organic carbon; DOC = dissolved organic carbon.
1. Introduction
Primary production measurement in marine waters is one of the most important tools to understand the ecosystem functioning and the transport of inorganic/organic matter through the food web. Direct measurement of the driving process, that is photosynthesis, in this dynamic system was first achieved using the Winkler technique for estimating dissolved oxygen concentrations, introduced by Gaarder and Gran in 1927 (Williams et al., 2002). Nowadays the light-dark bottle oxygen technique is considered not sensitive enough and the rate had poor accuracy and precision (Marra, 2002).
The introduction of the 14C technique changed the study of productivity (Marra, 2002). The year 1952 marked the end of a century-long struggle to develop a method to determine oceanic primary productivity with precision, accuracy and efficiency (Barber & Hilting, 2002). Thousands of measurements of marine phytoplankton productivity have been made at discrete locations throughout the world’s oceans since the introduction of the radiolabelled carbon uptake method in 1952 (Steemann-Nielsen, 1952). Although numerous, these discrete primary productivity measurements only provide information for infinitesimally small points over the oceans’ surfaces. Scaling these discrete measurements to global projections by means of satellite-based estimates requires mathematical models. Clearly, although much of the discrepancy between modelled and measured production results from limitations of the models, some of the disagreement is also due to methodological differences in 14C measurements and errors in the 14C data (Behrenfeld & Falkowski, 1997). To compare modelled primary production with that obtained
The 14C method is relatively simple. A known amount of 14C-CO2 is added to bottles containing the water samples and after an incubation time the organic carbon is frequently separated from the remaining inorganic carbon by filtration (Peterson, 1980). Several debates have arisen from the comparison of the results obtained with the 14C method and the Winkler method, the latter largely used for primary production estimates before 1952 (Peterson, 1980). In 1972 Schindler and co-workers modified the 14C method proposed 20 years earlier by Steemann-Nielsen. The authors described a method in which the samples were not filtered, in order to avoid the rupture of cells during filtration and loss during drying of filters that can still cause underestimation of primary production (Theodórsson & Bjarnason, 1975). The problems encountered in filtering and the somewhat arbitrary nature of the separation between dissolved and particulate matter led to the development of the acidification and bubbling procedure (Peterson, 1980). Unfortunately, this technique does not permit to assess the two fractions of primary production labelled with 14C-CO2: POC (Particulate Organic Carbon, corresponding to the 14C retained in the autotrophic cells) and DOC (Dissolved Organic Carbon, released from the autotrophic cells). Therefore this technique is not useful to understand the transformation of the organic matter along the trophic levels, in particular the link between autotrophic production and other levels of the marine food web. In most of the references regarding primary production in the water column, the method usually applied to stop the process and remove the excess labelled 14C was filtration and subsequent acidification with HCl (Table 1).
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Many authors use filters to estimate primary production, without considering the filtrated fraction, which corresponds to the exudate release, nor the total sample. Moreover, different HCl concentrations are used in the reported papers. When samples are filtered, filters can be wetted or rinsed with acid (Babin et al., 1994; Hewson et al., 2001) or, in more recent studies, exposed to concentrated acid fumes (Cermeño et al., 2006; Marañón et al., 2004; Teira et al., 2005). The added HCl concentration ranged from 0.1N (Mangoni et al., 2008) to 6N (Lewis & Smith, 1983; Pugnetti et al., 2005). Considering different acid volumes added to different sample volumes also the final acid concentrations were very diverse among the studies. Although the 14C technique has now been used in more than 35 000 oceanographic experiments (del Giorgio & Williams, 2005), from the review of the earlier and recent literature, we evidenced that different methods are still used to stop the photosynthetic activity, and that highly variable concentrations of HCl are added to remove the excess labelled bicarbonate which was not assimilated by the microalgae. In order to standardize this step of the 14C method, we designed a series of experiments, using both water samples and a mix culture of microalgal taxa which represent a coastal phytoplankton community. The first aim of our study was to define which HCl concentration, with an equal volume, among 0.1N, 0.2N, 0.5N, 1N, 2N and 5N was sufficient both to remove the excess labelled bicarbonate and to kill the cells but without damaging their structures with a consequent loss of assimilated 14C. Secondly, using both chlorophyll
2. Materials and methods
2.1. Water sampling
On 18th October 2006, 29th January 2007 and 4th September 2008 five litres of seawater was collected at 5 m depth by a Niskin bottle about 10 km far from the coast in the Gulf of Trieste (northern Adriatic Sea, Italy). In each cruise seawater temperature and chlorophyll
2.2. Experimental design
The next morning (10.00-11.00 a.m.) 1.320 l of sampled seawater was transferred to an Erlenmeyer flask and kept in the darkness for 30 minutes to reduce the photosynthetic activity. Subsequently, 99 µCi (3.66 MBq) of NaH14CO3 (DHI, Denmark) was added into the flask. The sample was gently stirred and then 6 subsamples of 220 ml were transferred to six Erlenmeyer flasks. Three of these were incubated at
2.3. Statistical analysis
Student's t-test was applied to test for significant differences between pairs of experiments considering different acid normalities for filters, filtrated and total samples. Only data obtained from the first reading was used for the t-test.
2.4. Effects of HCl on the planktonic microalgal community
In order to test if the added acid concentrations were sufficient to stop the microalgal photosynthetic activity, another experiment was carried out. A mix of six microalgae, representing the local phytoplankton community, was prepared, simulating the proportion among the groups which is characteristic of a resuspended coastal water mass. We chose:
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2.5. Recovery experiment
During microscopic observations we observed several shades of red autofluorescence. Since we could not be sure that a pale orange colour was a sign of a not viable cell, we carried out another test. We believed that some cells could be still viable after a mild acid treatment and thus able to photosynthesize, especially during the period elapsing between the addition of the acid and the addition of the scintillation liquid. We tried to demonstrate that after a mild acid treatment some microalgae were still photosynthetically active. Consequently, we restored their maintenance conditions looking at the microalgal growth after two weeks. Therefore, the residual aliquots (>4 ml) of a few selected treatments (without acid, 0.1N, 0.2N and 5N) from the first experiment (mix culture) were transferred into 100 ml Erlenmeyer flasks which were filled with F/2 medium to a final volume of 50 ml. The four flasks were kept at 15°C and 12:12 light:dark photoperiod until microscopic observations.
3. Results and discussion
3.1. In situ primary production as an analytical support to remote sensing
Frequently the problem with the model efficiency is data limited by the amount of representative
3.2. Different fixatives and HCl concentrations in phytoplankton primary production
Results of the phytoplankton primary production experiments are presented in separate figures as filter, filtrated and total fractions. In the graphs each bar represents one biological replicate, while standard deviations result from 2 scintillation counts (from the Quanta-Smart scintillation analyzer). In Figure 1 are shown filter DPM values of the three experiments. In the second experiment some problems took place during filtration, i.e. the process occurred too quickly probably due to not proper positioning of the following filters: 1L0.1N, 2L0.1N, 2L0.5N, 1L1N, 2L1N, 1L5N and 2L5N, therefore leading to an underestimate of DPM values.
In the same experiment we observed that DPM values of the first dark replicate were higher than those of the second dark replicate, which were in turn higher than the third one (except for the 5N treatment). Not considering the above mentioned replicates, we noted that both light and dark DPM values of the second experiment were higher than those of the first and third ones. Higher values were probably due to a higher chl
In Figure 2a-f are shown DPM values of filtrated samples of the three experiments. In the second experiment higher DPM values were registered due to the previously reported filtration issue. We infer that some of the labelled phytoplankton ended up in the vial placed underneath the corresponding filter without being retained by the filter. In fact there is a relationship between high DPM values of the filtrated samples and low DPM values of the corresponding filters (1L0.1N, 2L0.1N, 2L0.5N, 1L1N, 2L1N, 1L5N and 2L5N). Unlike the filters, where light values were 1 or 2 orders of magnitude higher than dark values, in the filtrated samples their difference was markedly lower. Since the volume of the filtrated sample was 5 times lower than that of the filter sample, DPM values obtained from the filtrated samples were generally 1 or 2 orders of magnitude lower than those obtained from the filters. When DPM values are very low, the biological variability among replicates is amplified. This variability was noted particularly in light and dark samples of the first experiment.
In Figure 3a-f are shown DPM values from total samples of the three experiments. Due to higher chl
While the average reduction of light DPM values between the 0.1N and 0.2N treatments was identical in the second and third experiments (6.62% and 6.61%, respectively), the percentage decrease in other pairs of treatments (0.1N
3.3. Statistical analysis
The t-test applied to dark samples of the three experiments highlighted only a few statistically significant differences between pairs of the tested acid normalities, e. g. for filters in the first experiment and the filtrated fraction in the last one, probably because DPM values were comparable to the background noise of the instrument. Except for the last experiment, performed in September 2008, the t-test did not show any differences between pairs of the tested acid normalities when performed on filtrated fractions, either (Table 3). In detail, in the first light-experiment the 2N acid treatment on filters was significantly different from the 0.1N, 0.2N and 0.5N ones, while the 5N treatment of the total light fraction was significantly diverse from almost all the other treatments. Similarly, in the same dark-experiments only the 5N acid treatment performed on filters was significantly different from the 0.1N, 0.2N and 0.5N ones. The most significant differences between pairs of acid treatments were obtained in the last experiment for the total light fraction.
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3.4. Effects of HCl on the planktonic microalgal community
The litmus test performed on the planktonic community revealed that the addition of 200 µl of HCl 0.1N was sufficient to decrease the pH value from 8 to 4 (Table 4, Fig. 4).
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The further increase of HCl normality gradually lowered the pH value down to zero. Since HCl is added to remove the excess labelled C, it is important to know the pH of the acidified sample. In fact, if HCl is not sufficiently concentrated, pH will not reach the value which is necessary to shift the equilibrium towards the CO2 fraction and consequently the removal of the labelled bicarbonate from the system. Only a pH < 5 ensures a complete shift towards the CO2 fraction (Libes, 1992). In water samples the 0.1N HCl treatment was enough to lower pH below this value. On the other hand, if the acid is too aggressive it can damage the cell membrane, causing the loss of an undetermined quantity of assimilated 14C and therefore leading to an underestimate of DPM values.
The macroscopic observation of the acidified mix culture samples revealed that while the control was brownish red coloured (mostly due to diatom pigments), 0.1N, 0.2N and 0.5N treatments were greenish. We hypothesize that in the first three mild treatments HCl degraded the accessory pigments, i.e. fucoxanthin, typical of diatoms and responsible for the brownish red colouring, but not chlorophyll
Another experiment on the planktonic community was performed where both motility of flagellates and of the tychopelagic diatom and chlorophyll
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Analysing samples in epifluorescence, we noticed that in the control all cells appeared red coloured. In the 0.1N treatment a different response was observed according to the taxon: the planktonic diatom, the naked dinoflagellate and the small phytoflagellate showed a less intense pigmentation (with respect to the control); half of the observed tychopelagic diatom cells was still red coloured, while the other half became faded. Only the benthic diatom and the thecate dinoflagellate were still bright red (Fig. 6).
In the 0.2N treatment the planktonic diatom and the small phytoflagellate did not show fluorescence any more, whereas the tychopelagic diatom and the naked dinoflagellate emitted a faint fluorescence.
Most of benthic diatom cells turned from red to orange, while only thecate dinoflagellate cells kept a bright red colour. Unlike the 0.2N treatment, the 0.5N sample presented naked dinoflagellate cells which lost their fluorescence and benthic diatom cells which were definitively orange. The only alteration in the next treatment (1N) was represented by tychopelagic diatoms which became pale. In the 2N treatment tychopelagic diatoms were not fluorescent any more, while thecate dinoflagellate cells turned their fluorescence from intense red to faded orange. In the last treatment (5N) benthic diatoms turned to faded orange, whereas thecate dinoflagellates emitted a faint fluorescence.
The microscopic observation of the recovered mix culture of the not acidified treatment revealed that some taxa were still alive and motile. The undetermined Cryptophycea was very active and abundant.
Considering the results of the litmus test and the microalgal viability test together, some considerations can be made. If the pH value is not sufficiently low, the inorganic 14C cannot be completely removed from the system and the cells remain still metabolically active. Consequently, the microalgae are able to continue both to uptake inorganic carbon and to consume the assimilated 14C during the period elapsing between the addition of the acid and the addition of the scintillation liquid. Moreover, if the pH value is sufficiently low to completely remove the inorganic 14C, but not to kill all the cells, primary production could be underestimated. In fact, with a complete depletion of inorganic carbon (pH <5), the cells cannot uptake it any more, but they could continue to consume the already assimilated 14C. This could occur treating the not filtered water samples with a mild acid.
Therefore we reckon that the acid concentration is crucial: only a sufficiently strong acid allows the complete removal of the inorganic carbon and at the same time the killing of all microalgal cells. In fact, when mild acid treatments were used, some taxa seemed to be not affected by the acid, as demonstrated by both our viability test and our recovery experiment. For example, the benthic diatom
4. Conclusion
The aim of this study was to define which HCl concentration, with an equal volume, among 0.1N, 0.2N, 0.5N, 1N, 2N and 5N (which correspond to final HCl concentrations of 0.004N-0.008N-0.02N-0.04N-0.08N-0.2N, respectively, for 5 ml filtrated and total samples) was sufficient both to remove the excess labelled bicarbonate and to kill the cells but without damaging their structures and therefore leading to loss of assimilated 14C. The litmus test demonstrated that the use of a mild acid (0.1N) does decrease the pH from 8 to 4. However, the results obtained from our viability test and recovery experiment suggest that some microalgal taxa in the not filtrated samples could remain still photosynthetically active after a too mild acid treatment. Therefore, the use of a slightly more concentrated acid (0.2N) is recommended. In our experiments this acid concentration led to on average 6% lower DPM values if compared to the 0.1N treatment, both for filters and total samples. Nevertheless, we believe that this small loss is acceptable in exchange for greater sureness of having killed the majority of the microalgal cells. In contrast, it is very likely that the 5N treatment damages the cell membrane with the consequent loss of assimilated 14C, leading to an underestimate of DPM values up to 31% for filters and 56% for total samples if compared to the mildest acid treatment. Our experiments were carried out using oligotrophic water (Fonda Umani et al., 2004) from the Gulf of Trieste. In the next future we are going to carry out another set of experiments on mesotrophic and eutrophic waters, applying the same chloridric acid concentrations, to test if similar results are achieved.
Acknowledgments
We thank Daniela Fornasaro and Benedetta Guardiani for helping us during the experiments. We are also grateful to Oriana Blasutto for water sampling and Massimo Celio for multiparameter probe data. Additional thanks go to Marina Cabrini and Paola Del Negro for the help they have provided in relation to this work. Finally, we thank Alfred Beran for providing us with microalgal cultures.
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