Upwelled plankton community modulates surface bloom succession and nutrient availability in a natural plankton assemblage

Paul, Allanah Joy; Bach, Lennart T.; Arístegui, Javier; von der Esch, Elisabeth; Hernández-Hernández, Nauzet; Piiparinen, Jonna; Ramajo, Laura; Spilling, Kristian; Riebesell, Ulf

doi:https://doi.org/10.5194/bg-2022-44

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https://doi.org/10.5194/bg-2022-44

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08 Mar 2022

Status: this preprint is currently under review for the journal BG.

Upwelled plankton community modulates surface bloom succession and nutrient availability in a natural plankton assemblage

Allanah Joy Paul¹, Lennart T. Bach², Javier Arístegui³, Elisabeth von der Esch¹, Nauzet Hernández-Hernández³, Jonna Piiparinen⁴, Laura Ramajo^5,6,7, Kristian Spilling^4,8, and Ulf Riebesell¹ Allanah Joy Paul et al.

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¹GEOMAR Helmholtz Centre for Ocean Research Kiel, Düsternbrooker Weg 20, 24105 Kiel, Germany
²Institute for Marine and Antarctic Studies, University of Tasmania, Hobart, Tasmania, Australia
³Instituto de Oceanografía y Cambio Global (IOCAG), Universidad de Las Palmas de Gran Canaria (ULPGC), Las Palmas, Spain
⁴Marine Research Centre, Finnish Environment Institute, Helsinki, Finland
⁵Center for Advanced Studies in Arid Zones (CEAZA), Coquimbo, Chile
⁶Departamento de Biología Marina, Facultad de Ciencias del Mar, Universidad Católica del Norte (UCN), Coquimbo, Chile
⁷Center for Climate and Resilience Research (CR)2, Santiago, Chile
⁸Centre for Coastal Research, University of Agder, Kristiansand, Norway

Received: 09 Feb 2022 – Discussion started: 08 Mar 2022

Abstract. Upwelling of nutrient rich waters into the sunlit surface layer of the ocean supports high primary productivity in Eastern Boundary Upwelling Systems (EBUS). However, subsurface waters not only contain macronutrients (N, P, Si) but also micronutrients, organic matter, and seed microbial communities that may modify the response to macronutrient inputs via upwelling. These additional factors are often neglected when investigating upwelling impacts on surface ocean productivity. Here, we investigated how different components of upwelled water (macronutrients, organic nutrients, seed communities) drive the response of surface plankton communities to upwelling in the Peruvian coastal zone. Results from our short term (10 days) study show that the most influential drivers in upwelled deep water are 1) the ratio of inorganic nutrients (NO_x : PO₄^3-) and 2) the microbial community present that can seed heterogeneity in phytoplankton succession and modify stoichiometry of residual inorganic nutrients after phytoplankton blooms. Hence, this study suggests that phytoplankton succession after upwelling is modified by factors other than the physical supply of inorganic nutrients. This would likely affect trophic transfer and overall productivity in these highly fertile marine ecosystems.

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Allanah Joy Paul et al.

Status: final response (author comments only)

RC1:
'Comment on bg-2022-44', Anonymous Referee #3, 18 Mar 2022
The manuscript shows a nice experiment planned to demonstrate the influence of the chemistry and biology of upwelled water on the development and later progress of phytoplankton blooms in coastal upwelling systems. In my opinion, the design of the experiment is correct, in which the simulation of the dilution caused by an upwelling episode stands out. However, upwelling most likely does not cause a full 1:1 mixing of upwelling water with surface water. Usually, upwelling pushes up, compresses the surface layer, and so supplies nutrients (and plankton) by diffusion and turbulent mixing at different intensities.

The conclusions are correct, although expected. The first conclusion is the best known. That is, nutrient supply to surface water with low nutrient concentrations induces phytoplankton blooms, mainly diatoms. The second is somewhat new, but not strange. It is reasonable to expect that plankton populations reaching the surface with upwelled waters modulate the bloom and its later evolution. This experiment clearly demonstrates that this happens. However, a better characterization of the species and/or genera of phytoplankton involved is lacking. The flow cytometer has only allowed the characterization of Synechococcus. For the rest of the community there was only a proxy of its size with very low detection of microphytoplankton. In addition, chlorophyll was not fractionated. This information is especially important in the post-bloom, when divergence between treatments and the variability within treatments is more evident. However, there is also variability among treatments during the bloom, as inferred from the differences in chlorophyll concentration on day 4 (Fig. 3A) and in the different abundances of nanoplankton (Fig. 5D, E, F). On the other hand, the results reporting significant silicate drawdown in the HN biology treatment point to the importance of diatoms, which could be different from those found in other treatments, including the LN biology treatment. Microphytoplankton (mainly diatoms) are likely causing the divergence observed in both bloom and post-bloom.

Although the introduction and the discussion read well, this is not the case for the results. In my opinion, this section, of great importance to support the conclusions, is written in a cumbersome way. It is necessary to read it several times and with enough attention to catch the information. Figures are not always properly cited, nor is supplementary material. There are tables in the supplementary material that are not cited in the text.

In my opinion, this results section could be improved to remove weaknesses and make the manuscript more attractive to potential readers. The manuscript will probably improve by focusing the description of results on those relevant to the conclusions and ignoring those with low contribution to the two main conclusions.

Despite the lack of information on phytoplankton species composition, which in my opinion represents the greatest weakness of the manuscript, the design of the experiment and the difficulty of its execution, lead me to recommend the publication of the manuscript with major revisions.

Specific comments

Introduction

Line 29. …is considered the most productive… from where? Maybe …”the most productive upwelling region” or something similar

Lines 66-68. The Peruvian productivity paradox is a common paradox to all upwelling systems. With strong upwelling, chlorophyll concentration is low because surface water is recently upwelled water with high nutrient concentrations. Chlorophyll concentration increases when upwelling relaxes. This time lag also has translation into spatial heterogeneity. Chlorophyll concentration is low (few phytoplankton) in the upwelling center where there is deep water recently upwelled. High chlorophyll levels can be found in the surroundings.

Materials and methods

Line 105. According to Figure 1, the range of 15 m was only at station A, at station B it was 5 m.

Line 109. …collected from the mesocosms

Lines 129-131. The last sentence reads, “Both the surface (mesocosms) and treatment water (deep water) were filtered… However, the deep water added to the two biology treatments was unfiltered.

Line 135 …were set to the same as in the organic…

Line 173. (picoeukaryotes, , small microphytoplankton, large microphytoplankton). It may be appropriate to add a few words here to inform that microphytoplankton is not well estimated by this technique, although it is recognized in the legend of figure S2.

Results

Initial conditions (Day 1)

Lines 256-257. If referring to all nutrients, Fig. 3B and C and Table S1 should be cited. If only nitrate is referred to, Fig. 3B should be cited.

Lines 262-265. Fig. 4E should be cited when discussing a254. For E2:E3 it should be Fig 4F. Add Table S1 to Fig 3F when LAP activity is discussed; the slightly higher activity is better seen in the table than in the figure.

Lines 266-271. Table S1 should be mentioned when commenting about the phytoplankton community. The same table can be mentioned for Fv/Fm, Fig. 4D is the figure.

Line 274. …between Day 3 and 5. Better between Day 3 and 6 (Fig. 3D).

It is difficult to follow the chlorophyll in this paragraph, it would be better to specify something else, for example: Peak Chl a concentrations of up to 12 µg L-1 (HN organic) and ~6 µg L-1 (LN inorganic and biology). According to figure 4A, there are differences between various treatments on this day 4.

It is difficult to follow this about the ratio DIN drawdown to maximum Chla accumulation. This ratio was higher in LN only for the case of organic treatment (Fig. 4A). I think the next paragraph about higher recycling of N or highest N utilization efficiency under low nitrate needs further explanation. How this higher N recycling or N utilization efficiency deduces from a lower DIN ratio drawdown to Chla accumulation? It seems too risky to attribute these differences in the ratio only to N. Variations in the ratio may also be due to different cell concentrations of chlorophyll. Mixotrophic behavior can also affect this ratio. The ratio changes through changes in N, changes in chlorophyll, or in both. Here phytoplankton composition could provide additional information.

Lines 291-292. The last sentence indicating that the initial concentration of DIN was 3 times higher in HN than in LN can be deleted. It was reported at the beginning of the results.

Line 322-323. I understand the association between higher silicate drawdown and higher chlorophyll concentration, but not with nanoplankton abundance. There is no information on the species that are in the nanoplankton fraction. On the other hand, the increase in chlorophyll could well occur in micro diatoms. Maybe the sentence could write like this:

The highest Si(OH)4 and phosphate drawdown, and consequently Chl a concentration was observed in one replicate. This replicate also showed highest nanophytoplankton abundances (Fig. 5B).

Fig. 5. I think the symbols on the panels do not correspond to the ones on the labels, where they are all circles.

Discussion

Line 370-372. I think this sentence about bottom-up and grazing control is missing something.

Line 420-422. Silicic acid consumption could well have occurred by micro-sized diatoms. It is difficult to conceive that all or nearly all of the nanophytoplankton were diatoms. Usually, there are many flagellates in this fraction.

Lines 429-430. Diatoms were not analyzed and, therefore, it cannot be confirmed that the different behavior of the two treatments was due to the different response of the diatoms and the different seed population. What can be said is that the different behavior of the two treatments could be attributed to a different response of the diatoms and probably also to differences in the seed population.

Line 490-491. The highest silicate uptake only occurred in a biology treatment, in the HN biology. In the LN biology it did not occur (Fig. 3D).
Citation: https://doi.org/10.5194/bg-2022-44-RC1
RC2: 'Comment on bg-2022-44', Anonymous Referee #4, 24 Jun 2022

Review Paul et al « Upwelled plankton community modulates surface bloom succession and nutrient availability…… » submitted to Biogeosciences

General comments.

This ms describes the time evolution of nutrients, chlorophyll, flow cytometric groups, chromophoric dissolved organic matter in experimental 15L bags during 10 days after mixing surface sea water with an equivalent volume of ‘deep’ water under different conditions (nutrient rich, nutrient poor, filtered (with natural organic matter and nutrients but no organisms) or not (with natural nutrients, organic matter and deep microorganisms). A nutrient control was set by mixing surface sea water with filtered surface sea water and adding nitrate and phosphate. The design of the experiment was set to explore not only the effect of nutrients on the sea surface nutrient consumption and phytoplankton dynamics and composition under the influence of upwelling episodes, but also that of nutrients + organic matter, and that of nutrients + organic matter + natural deep communities upwelled from the ‘deep waters’.

If the objectives of the experiment are clear, a huge work and many parameters shown, an interesting introduction and discussion, the design and the results are very complicated to follow. I suggest to the authors to simplify denominations/definitions of the different combinations in the bags and to rewrite the m&M and the result section to give it clearer. In addition, I was left a bit frustrated as:

For a biogeochemist point of view, we have no information on ammonium concentrations. Regenerations sources could have been high in the bags, particularly considering the high concentrations of nitrate added: around 8 and 3 µM in HN and LN, respectively. This information should have been pertinent to state nitrogen limitation status and examine N/P ratios (and for this reason, you should use the term NOx instead of DIN for the whole text, as it refers only to the sum nitrite + nitrate)

For a microbiologist point of view, again, we have only partial information:

First on the microphytoplankton response: There is discussion on a diatom response, but we have not any information on the taxomomic composition of diatom communities, even using proxies that could be have been given for instance by size fractionation of chlorophyll. Indeed, flow cytometry analysis only allows small size class to be counted. There was only few information given on a “chain group”, on a “small microphytoplankton group”, and on a “large phytoplankton group” which was not statistically represented in the flow cytometry analysis (Fig. S2), and furthermore the time evolution of abundances in these groups is not plotted (just initial conditions, table S1) or statistics (table 2).

Second, there is no information on heterotrophs. There is discussion on the potential role of heterotrophic bacteria on the degradation of organic matter, and on the the phytoplankton regulating process during its decay period. But not any data is available on heterotrophic bacterial abundances and/or on the grazer community composition, or virus abundances.

The choice of deep samples at 90-105 m at station A for HN experiment, and 40-45m at station B for LN experiment is crucial to compare experiments. Clearly the vertical distributions of physical properties, nutrients, abundances of flow cytometric groups and organic matter properties at these 2 stations, could have been helpful for the readers, particularly those not familiar with the biogeochemical context in this area, to situate these “deep” water masses in the frame of depths of nutriclines, OMZ, deep chlorophyll maxima, euphotic zone depth etc… Were these depths taken out of the euphotic zone? out of the OMZ?, Were they constituted only by heterotrophs or were there also some phytoplankton?

And Lastly, why taking waters from the mesocosm experiment instead to in situ surface waters close to the mesocosm? Why day 20?

For all these reasons, I would recommend the publication of the ms with major revisisons.

Specific comments

Line 109. Specify even of you refer to Bach et al what was the status of phytoplankton on the day 20 of the mesocosm study: steady state? exponentially growing? decaying?

Line 119. The authors should moderate this sentence “… no TM or DOM measurements were made… these were assumed to be different…. ”

Figure 2. station A and station B should be removed in the lines “inorganic”, somewhere it should be drawn that nitrates and phosphate were added. In the legend for memory it should be reminded that “unfiltered” is in fact < 64 µm and the “filtered” is < 0.1 µm.

Line 135 for the “inorganic “treatment why not also adding Silicates to get similar changes for N/Si ratios?

Line 138, 140. Be more precise and everywhere add the porosity of the filtration.

Line 172. As the samples were not fixed, how were stored samples between sampling and analysis? was there a long delay between the first and the last sample analyzed?

Line171-180. The authors should add more details on the set up of the flow cytometer (and/or on the legend of Figure S2): debit and duration of the analysis, i.e. the volume analyzed), also did you add beads to set limits between the different size classes and how this differentiation was done. Add information on wavelengths of the different windows (FL3A, FL2H, FL4H) that could be indicated on the legend of the Figure S2. In this figure some cytograms were excluding some populations and other not? explain more in the legend. As demo, a plot FL2 / FL3 would have been useful too.

Could you really consider counts of micro-II as significant? we just have initial conditions in terms of percentage on table S1, but no idea of any abundance,

For a paper dealing on the potential effect of seeding microorganisms with upwelled waters, I find rather strange to get so much groups counted with the cytometer and having finally only a plot of the evolution of and nanophytoplankton.

Were abundances detectable?

Line 190. Were all the FV/Fm measurements done on the same time of the day?

Paragraph 2.3 Is there any information on the distribution of heterotrophs? heterotrophic bacteria? Heterotrophic nanoflagellates? ciliates? the filtration on < 64 µm could have a cascading high effect on ciliates, which becomes the top predator in the bags.

Line 229. Please explain better for non-specialists: “the contrast matrix… the organic treatment was used as the control for the linear mixed model analysis”

Line 256. refer also to Table S1

Line 259. refer a254 with Figure 4e, and on line 262 E2:E3 ratio with fig 4f

Line 264. “surface water…. nitrogen depletion”. Initial concentrations in NOx in “inorganic” (2.07) versus those in organic LN (2.49 and 3.17 (organic LN) were not so different and where all evolving the same way considering nitrate (Figure 3b) or LAP activity (Figure 3F)

Line 266. please explain in the Material and method in 2.3 section clearly how is calculated the “relative contribution of each group to chlorophyll a fluorescence in flow cytometric analysis” Also if it is this parameter used on table S1 when reporting percentages of the different flow cytometric groups, and not simply relative abundances, it should be explained in the legend.

Line 268. Change formulation “R statistic” here and in Table 2 Not the software, but the type of test should be indicated.

Line 273. It is not visible in Fig 3B plot, were the limits of detection for nitrate (0.123 µm as stated in the methods) reached in all experiments?

Line 282. Is there an error here? I would rather write this sentence like this: “DIP is more consumed relative to DIN in LN treatments compared to HN treatments ….”

Line 283-284. Is there an error here? rather it should be 9.82 for HN and 6.25 for LN?

Line 285. “… where initial N was lowest”. No, initial N was the lowest in the inorganic treatment.

Line 289. “higher recycling… “. Ammonium concentrations were not available? Probably like DIP it was produced by regeneration between days 6 and 8, see lines 325-326

Line 289. “ .. or highest N utilization efficiency under low nitrate” There is another hypothesis, a higher top-down control of phytoplankton by grazers under high nitrate.

Lines 291-292. This sentence on initial nitrate nitrite concentrations should be cited at the beginning of section 3.1

Figure 3. Why plotting silicate drawdown when absolute concentrations are plotted for Chl, DIN and DIP? For plots based on ‘deltas’ like figure 3D and 3E the authors should explain in the legend if the difference is always made with T1 concentrations

Figure 4A. For the legend, indicate how DIN was calculated. Was it DIN at T1 minus DIN at the time of max chlorophyll, i.e. T4 for all samples except T3 for LN organic?

Figure 4B. Again it is unclear if the difference is made as concentrations at day 6 minus concentrations at day 10 of these box plots are simply the means of the data presented figure 3E for the period T6 to T10. Be clearer in the legend. Is seems that here are presented the distribution of the 20 data (T6 to T10 time points x quadruplicates bags).

Line 324. Refer to fig 5E

Line 330. The last sentence with infos on initial conditions should be cited in section 3.1

Table 2. Modify “R statistic”, indicate in the legend what were the bloom and post bloom periods considered for the tests. For the relative contribution, in percentages, I don’t understand to what they refer, as the sum of contribution of each group does not make 100%.

Line 369. It is up to 12 µg/l as seen from the figure 3A

Line 372. “... than any impact of grazing”: But some grazers were present in the surface water taken in the minicosms. Furthermore, this surface water was filtered through 64 µm, and consequently with no top predators, all microzooplankton (heterotrophic ciliates) could have been rapidly growing in response to the increase of pico nano and small microphytoplankton. Note also that his sentence lines 370-372 has no verb.

Line 372-374. Rather, I would imagine than heterotrophic bacteria would find more favorable growth conditions with surface water mixed with deep filtered sea water, as the surface heterotrophic bacteria are diluted in deep water by a factor 2 as well as their grazers, and thus have less predatory control, together with more access to nutrients and DOM provided by the deep waters.

Is there any information on abundances of heterotrophs? heterotrophic bacteria? flagellates? ciliates?

Line 385. The noticeable net increase of DIP in experiments between T6 and T8 suggests that ammonia could have been also regenerated through grazing processes during that period. This increase of DIP about 0.1 to 0.3 µmole/l, based on a N/P ratio of 16, could signify that as much as 1.6 to 4.8 µM of ammonia could have been regenerated, even based on a delta DIN/delta DIP of about 6, this give up to about 2 µmole/l ammonia regenerated.

Line 393. The authors should cite the initial DIN/DIP ratios here, and write that DIP was never depleted in the experiments.

Line 396. Without ammonia measurements, it is difficult to speculate on N regeneration. However, if abundances of heterotrophic prokaryotes are available, I suggest to calculate per cell LAP activities.

Line 401 “LAP was higher…”. Before comparison with other studies the concentration of leu-AMC used by other authors should be verified as it influences rates. The concentration added here (500 µM) is high. Mabmig et al used 200 µM.

Line 428. “Irradiance levels increased upon incubation”. Were the levels of irradiance in bags higher than in the surface mesocosms?

Line 440 “viral presence”. Because the authors made a 0.1 µm filtrations the ratio viruses to their host is very high in initial conditions., could it be in the favor of viral lysis?

Line 433. “… rather then the manipulated deep water”. It would have been interesting to have initial compositions of populations included in the two “deep waters” used in this experiment.

Line 455. “...and higher post-bloom Chla concentrations were sustained in this treatment”. Yes, but in the “inorganic” too, so the source of the variability is not only due to the variability of responses of the seeded communities, those of surface too.

Line 480. Sentence unclear, does the term “that” refer to physical factors? If yes do you discuss about the horizontal mixing by showing the example of tidal mixing? If yes write it.

Citation: https://doi.org/10.5194/bg-2022-44-RC2

Allanah Joy Paul et al.

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Short summary

We investigated how different deep water chemistry and biology modulate the response of surface plankton communities to upwelling in the Peruvian coastal zone. Our results show that the most influential drivers were the ratio of inorganic nutrients (NO_x : PO₄^3-) and the microbial community present in deep water. These led to unexpected responses in the phytoplankton assemblage that could not be predicted by the amount of inorganic nutrients alone.


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