# Dissolved organic matter released by two marine heterotrophic bacterial strains and its bioavailability for natural prokaryotic communities

Eva Ortega-Retuerta, Quentin Devresse, Jocelyne Caparros, Barbara Marie, Olivier Crispi, Philippe Catala, Fabien Joux, Ingrid Obernosterer

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## To cite this version:

Eva Ortega-Retuerta, Quentin Devresse, Jocelyne Caparros, Barbara Marie, Olivier Crispi, et al. "Dissolved organic matter released by two marine heterotrophic bacterial strains and its bioavailability for natural prokaryotic communities." Environmental Microbiology, 2021, 23 (3), pp.1363-1378. 10.1111/1462-2920.15306. hal-03007145

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Submitted on 16 Nov 2020

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Distributed under a Creative Commons Attribution 4.0 International License
1 Dissolved organic matter released by two marine heterotrophic bacterial strains

2 and its bioavailability for natural prokaryotic communities

3 Eva Ortega-Retuerta, Quentin Devresse*, Jocelyne Caparros, Barbara Marie, Olivier

4 Crispi, Philippe Catala, Fabien Joux, Ingrid Obernosterer

5 CNRS/Sorbonne Université, UMR7621 Laboratoire d'Océanographie Microbienne, Banyuls sur Mer, France

6 * Present address: Geomar Helmholtz Centre for Ocean Research Kiel, Germany

7

8 Running title: Bacterial DOM release and bioavailability
10 **Originality-Significance Statement**

11 The role of heterotrophic prokaryotes as organic matter sources in the ocean is
12 increasingly receiving attention by the scientific community in view of its importance
13 for major biogeochemical fluxes, such as carbon sequestration via the “microbial carbon
14 pump”. Here we present novel data on dissolved organic matter (DOM) production by
15 bacterial strains and how this bacterial-derived DOM is degraded by in situ prokaryotic
16 communities and how bacterial DOM shapes microbial metabolism and diversity. We
17 demonstrate that DOM quality produced by bacteria depends on their taxonomy, which
18 translates into different degradation mechanisms and the growth of different microbial
19 communities. Even though a high amount of bacterial derived DOM can be degraded
20 within days, this DOM source would lower the prokaryotic growth efficiency of
21 degrading microbial communities compared to in situ DOM.

22
23

## 24 Summary

Marine heterotrophic prokaryotes (HP) play a key role in organic matter processing in oceans; however, the view of HP as dissolved organic matter (DOM) sources remains underexplored. In this study, we quantified and optically characterized the DOM produced by two single marine bacterial strains. We then tested the availability of these DOM sources to in situ Mediterranean Sea HP communities. Two bacterial strains were used: *Photobacterium angustum* (a copiotrophic gammaproteobacterium) and *Sphingopyxis alaskensis* (an oligotrophic alphaproteobacterium). When cultivated on glucose as the sole carbon source, the two strains released from 7 to 23% of initial glucose as bacterial derived DOM (B-DOM), the quality of which (as enrichment in humic or protein-like substances) differed between strains. B-DOM promoted significant growth and carbon consumption of natural HP communities, suggesting that it was partly labile. However, B-DOM consistently promoted lower prokaryotic growth efficiencies than in situ DOM. In addition, B-DOM changed HP exoenzymatic activities enhancing aminopeptidase activity when degrading *P. angustum* DOM, and alkaline phosphatase activity when using *S. alaskensis* DOM, and promoted differences in HP diversity and composition. DOM produced by HP affects in situ prokaryotic metabolism and diversity, thus changing the pathways for DOM cycling (e.g. respiration over biomass production) in the ocean.

43

Keywords: microbial carbon pump, dissolved organic matter, bioavailability, heterotrophic prokaryotes, prokaryotic diversity.
46

## 47 Introduction

Heterotrophic prokaryotes (HP, bacteria and archaea) are key players in element cycling in the ocean. Roughly 50% of carbon that is fixed by primary production in the ocean is passing through HP (Azam et al., 1983), which can be respired into CO₂ (HP respiration) or used to build up new biomass (HP production). The balance between these two processes, expressed as prokaryotic growth efficiency, is a key parameter to estimate carbon fluxes in the ocean and predict or model its changes (delGiorgio and Cole, 1998). However, it has been demonstrated that HP also release carbon in the form of dissolved organic matter (DOM). Pioneering studies showed that HP growing on simple substrates (i.e. glucose) could produce a myriad of different DOM compounds (Stoderegger and Herndl, 1998; Ogawa et al., 2001; Kawasaki and Benner, 2006). Since then, numerous studies have attempted to characterize HP-derived DOM using analytical techniques such as ultra-high-resolution Fourier transform-ion cyclotron resonance-mass spectrometry (FT-ICR MS), high field nuclear magnetic resonance (NMR) or 3D fluorescence spectroscopy, demonstrating that HP successively process labile DOM and transform it into refractory DOM in an amount enough to sustain the whole oceanic refractory DOM pool (Catalá et al., 2015; Lechtenfeld et al., 2015; Osterholz et al., 2015). This process, termed the microbial carbon pump (MCP, Jiao et al., 2010) is an important carbon sequestration mechanism. It has been estimated that carbon stored through the MCP accounts for 0.4% of primary production on a global scale; this is in the same range as carbon stored through sinking particulate organic matter (0.6 to 1.3% of primary production (Legendre et al., 2015)).

HP release DOM as metabolic by-products or as a consequence of the uncoupling of anabolic and catabolic processes (Carlson et al., 2007). Since HP in the ocean exhibit a
wide range of metabolisms, DOM compounds released to the media are expected to differ among HP taxa, as demonstrated previously using experimental incubations (Shimotori et al., 2012; Noriega-Ortega et al., 2019). HP also actively release DOM compounds for different processes such as polymer enzymatic cleaving (Smith et al., 1992), nutrient acquisition (e.g. siderophores for iron uptake (Andrews et al., 2003), cell-cell communication (e.g. homoseryl lactones for quorum sensing (Gram et al., 2002)) or toxins (Christie-Oleza et al., 2012). Thus, environmental variables could shape the quantity and quality of the DOM released, affecting DOM fluxes. Finally, DOM degradation in the ocean, including HP-derived DOM, could depend on the taxonomic composition of the DOM processing community (Teeling et al., 2012). To better understand and predict changes in carbon sequestration through the microbial carbon pump, accurate information about DOM production by heterotrophic prokaryotes and its cycling in the ocean is a priority. How HP-derived DOM is degraded and cycled, and which fraction of it would be respired to CO₂ or would enter the oceanic refractory DOM pool, are questions that remain to be resolved.

In this study, we aimed to evaluate the availability of DOM released by two single bacterial strains as a substrate for Mediterranean Sea heterotrophic prokaryotic communities. Our working hypotheses were (1) DOM quality differs among bacterial strains and (2) these differences in quality promote the growth of different prokaryotic communities, translated into different DOM degradation mechanisms.

To test these hypotheses, two step experiments were conducted: First, two single bacterial strains with contrasting lifestyles and from different taxonomical classes (*Photobacterium angustum* and *Sphingopyxis alaskensis*) were axenically grown in minimum media with glucose as the only carbon source, and cultures were harvested at early stationary phase where DOM was extracted, quantified and characterized. Second,
96 DOM derived from the strains was used as substrate for biodegradation experiments,
97 where in situ prokaryotic communities from the Mediterranean Sea were grown either
98 on in situ DOM or on the DOM produced by the two strains (Table 1, see experimental
99 procedures). In these experiments, changes in heterotrophic prokaryotic abundance were
100 followed together with changes in carbon and nutrients, exoenzyme activities and
101 prokaryotic community composition.
102
103 **Results**

104 **Bacterial strain growth: Glucose and nutrient consumption; BDOM production**

105 The two single strains used in the present study to produce bacterial-derived DOM
106 exhibited contrasting patterns both in their growth rates and carrying capacities (Table
107 2, Suppl. Fig. 1): *Photobacterium angustum* grew fast (growth rates 1.1-1.5 d⁻¹), this is,
108 once the exponential growth started, the stationary phase was reached about 12h later,
109 with a cell abundance of around 5 x10⁶ cell mL⁻¹ (Suppl. Fig. 1). Conversely,
110 *Sphingopyxis alaskensis* grew slower (growth rates of 0.07-0.23 d⁻¹), with a longer lag-
111 phase (3 to 5 days) but higher cell abundance (12 to 17 x10⁶ cell mL⁻¹, Suppl. Fig. 1)
112 when the stationary phase was achieved.

113 When cells and DOM were harvested, 12 to 17 µM of carbon as glucose was remaining
114 in the cultures (6-10% of initial glucose concentrations, Fig. 1). However, there was 42
115 to 73 µM of DOC remaining (Fig. 1). Comparing the decreases in glucose and in DOC
116 at the beginning and at the end of the incubations (see Experimental Procedures), we
117 estimated the amount of DOC that was produced by the bacterial strains. This DOC
118 production was variable, ranging from 6.7% (S. alaskensis, March 2018) to 22.8% (P.
119 angustum, March 2018) of the initial DOC. Bacteria passing through the filters
120 represented less than 1% of cell abundances, and could only contribute to a tiny part of
121 the DOC measured at the end of the cultures. No clear differences between strains could
122 be observed in the %DOC produced (Table 2). However, the two strains produced
123 DOM of different quality. DOM produced by P. angustum was relatively more enriched
124 in protein-like compounds (FDOM Peak T after Coble (1996), 34.3 x10⁻³ R.U. on
125 average) as compared to DOM from S. alaskensis (8.9 x10⁻³ R.U. on average). By
126 contrast, all humic-like FDOM peaks were higher for S. alaskensis, in particular the M
127 and A peaks that were twice as high as for P. angustum (Table 2). Regarding inorganic
128 nutrients, both strains used preferentially NH₄ over NO₃, but significant amounts of
129 NH₄, PO₄ and NO₃ remained in the cultures at the end of the incubations
130 (Supplementary Table 1), indicating that carbon was the limiting substrate for the
131 bacterial growth.

132 **Availability of bacterial-derived DOM for strain-derived natural heterotrophic**
133 **prokaryotic communities**

134 *Biodegradation experiments: Changes in HP abundance, DOC and growth efficiency*

135 DOM originating from the bacterial strains promoted remarkable HP growth in all
136 biodegradation experiments (Fig. 2), with significant differences over time and between
137 treatments in all three experiments (repeated measures ANOVA, p < 0.05, Suppl. Table
138 2). In the open sea-March biodegradation (Table 1), DOM from *P. angustum* supported
139 a higher HP growth for surface and deep HP communities than their respective
140 treatments with in situ DOM (Fig. 2a), although only significant for surface
141 communities (Suppl. Table 3). In the coastal sea-April biodegradation experiment,
142 differences in the HP abundances among treatments were only significant at certain
143 timepoints (Fig. 2b, Suppl. Table 3). In the coastal sea-December biodegradation
144 experiment, as in the open sea-March one, the treatments with bacterial DOM promoted
145 significantly higher HP growth than *in situ* DOM (Fig. 2c, Suppl. Table 3). In the
146 coastal sea-December experiment we included a treatment with glucose (20 µmol C L⁻¹), but although HP abundance was higher than that in the surface DOM treatment, it
147 was lower than HP abundances in the *S. alaskensis* DOM treatment (Suppl. Table 3).
148 We did not find significant differences in HP growth between treatments using *P. angustum* vs. *S. alaskensis* DOM (Suppl. Table 3).
149
150
We performed controls in the degradation experiments, consisting of DOM from each strain, with N and P additions similar to the treatments, but without inoculating with in situ HP communities. The purpose of these controls was to check for potential growth of the minimal amount of *P. angustum* and *S. alaskensis* cells that remained in the filtrates. Little cell growth was observed in these controls (Fig. 2a-c).

At the onset of the three biodegradation experiments, the DOC concentration was slightly different between treatments (Table 3). However, the amount of DOC consumed per hour (during the total incubation) was consistently higher in the treatments with strain derived DOM, except for coastal sea-April experiment. Biodegradation experiment and *S. alaskensis* DOM (Table 3). Average DOC consumption in the incubations with strain DOM (both *P. angustum* and *S. alaskensis* DOM) was 0.30 µmol L⁻¹ h⁻¹, while in the treatments with in situ DOM, either deep or surface, the average was 0.08 µmol L⁻¹ h⁻¹. In the experiments with glucose additions the DOC consumption was on average 0.18 µmol L⁻¹ h⁻¹. When normalizing ΔDOC by the initial DOC concentration in each treatment, these patterns were even more visible: 51% on average of the strain-derived DOC was degraded during the incubations, compared to 13.6% of in situ DOC (Table 3).

We further wanted to elucidate whether differences in HP growth were due to different initial amounts of DOC or to differences in its lability. For that purpose, we compared HP biomass changes with initial DOC concentrations in every treatment (Table 3). In contrast to total DOC utilization, we could not find consistent patterns in the relative amount of initial DOC incorporated into HP biomass (Table 3). The %DOC invested into prokaryotic biomass was lower (open sea-March), similar (coastal sea-April) or higher (coastal sea-December) than in the in situ DOM (ndom) treatments. Comparing the treatments with *P. angustum* (pdom) or *S. alaskensis* (sdom), the changes in HP
biomass compared to initial DOC were never significantly different. Interestingly, prokaryotic growth efficiency (PGE) was consistently lower in treatments with strain-derived DOM than in the treatments with in situ DOM in all biodegradation experiments by a mean factor of 3.5 (range 1.3-7.0) (Fig. 3).

## Biodegradation experiments: Changes in FDOM and inorganic nutrients

The different FDOM peaks did not decrease at the same level in every incubation (Table 3): The peak T (surrogate of protein-like substances) tended to decrease up to a 70%, while the FDOM at peak C (surrogate of humic-like compounds) tended to increase in almost every experiment and treatment (Table 3). Humic-like peaks A and M were more variable among experiments and treatments.

The inorganic nutrient concentrations, particularly those of NO₃⁻, were also higher in the treatments with bacterial-derived DOM as compared to the in situ DOM, since *P. angustum* and *S. alaskensis* did not consume all inorganic nutrients during the previous strain incubations and equivalent amounts of all inorganic nutrients were added to each treatment of the biodegradation experiments. At the end of the incubations, substantial amounts of all inorganic nutrients remained in all treatments (Supplementary Table 1).

## Coastal biodegradation experiments: Changes in exoenzymatic activity

Differences in the predominant exoenzymatic activities could be observed between treatments in Coastal sea-April and December experiments. In May, relatively high alpha- and beta- glucosidase production rates were observed in the treatment with surface DOM (Fig. 4 a,b) compared to the bacterial DOM treatments, but this result was not visible in December (Fig. 4 e,f). Remarkably, in both biodegradation experiments *P. angustum* DOM induced high rates of leucine aminopeptidase (Fig. 4 c and g), while the *S. alaskensis* DOM always induced high rates of alkaline phosphatase (Fig. 4 d and h).
200 Coastal-December Biodegradation experiment: Changes in prokaryotic community
201 composition

202 Changes in HP diversity and community composition along the incubations were
detected in all treatments and replicates of Coastal sea-December experiment, the only
203 experiment where these samples were taken. A decrease in diversity between the onset
204 and end of the incubations, expressed as richness, evenness or Shannon diversity, was
205 detected in all treatments. Significant differences between treatments were observed at
206 tf (ANOVA, p < 0.001 for all richness, evenness and Shannon). HP communities
207 growing on surface DOM had higher richness, evenness and Shannon than the
208 communities growing on bacterial-derived DOM (Table 4).
209

210 At the onset of the experiments, the HP community was distributed among different
211 orders, with Alphaproteobacteria (groups of SAR11, Rhodobacterales and
212 Rhodospirilales), SAR86 (Gammaproteobacteria) and Flavobacteriales as the most
213 abundant groups (Fig. 5). At the end of the incubations, differences between treatments
214 could be detected (ANOSIM test, p < 0.05): The surface DOM treatment was
215 dominated by Alteromonadales, Flavobacteriales and SAR86, the surface DOM
216 +glucose was dominated by Vibrionales and Flavobacteriales (except for replicate 3
217 where Oceanospirilales were comparatively more abundant), the *P. angustum* DOM
218 treatments were dominated by Alteromonadales, Rhodobacterales, Flavobacteriales (but
219 replicate 2 showed a potential contamination by Campylobacter) and the *S. alaskensis*
220 DOM treatment was dominated by Vibrionales, with more than half of the sequences
221 corresponding to this order, but also Rhodospirilales. A MDS ordination based on Bray
222 Curtis similarities showed separation among treatments with replicates grouping
223 together, illustrating the significant differences (ANOSIM test, p < 0.05) in HP structure
224 (Fig. 5).
225 **Discussion**

226 **Comparison of DOC release rates among studies (strains vs in situ communities)**

227 The bacterial DOC release rates reported here (7 to 23% of initial DOC, average 12%) 228 are within those previously published for mixed/natural communities (3.5-22%) but 229 higher than the four published studies using single bacterial strains (3.7% on average, 230 Table 5). All previously published studies using single strains used 5 to 350 times 231 higher initial DOC concentrations than in the present study, and some only quantified 232 the DOM retained by solid-phase cartridges (Romano et al., 2014) or only the DOM 233 released as extracellular polysaccharide (Zhang et al., 2015), making the values lower.

234 The only study measuring DOC production by Archaea through chemoautotrophy also 235 reported DOM release within our ranges when looking at DOM produced vs. biomass 236 produced (Bayer et al., 2019). We could expect that the net DOM production by natural 237 communities would be lower than the ones recorded using single strains, since DOM 238 produced by some HP groups could be consumed by others. However, our DOM 239 production values were within the range of previous studies using mixed communities 240 (i.e., 3.5-22%, Table 5). The DOM production values obtained in our strain incubations 241 could be conservative estimates. On one hand, the method used to quantify glucose 242 (Myklestad et al. 1997) also determines other dissolved monosaccharides. Although it is 243 very unlikely that the strains would have released these compounds without making use 244 of them, their potential presence would lead to an underestimation of the DOM release 245 rates in our experiments. On the other hand, HP release DOM not only as metabolic by- 246 products, but also with functional roles such as cell-signaling (Gram et al., 2002) or 247 particle degradation (Smith et al., 1992) among many others, processes that would not 248 be happening in our simple glucose-amended monocultures, so DOM production in situ 249 by HP would likely be higher than the calculated in our study. For instance, substrate
quality is likely crucial determining the quantity and quality of DOM produced: Ogawa et. al. (2001) and Wienhausen et. al. (2017) showed higher release of dissolved amino acids by heterotrophic bacterial strains when growing on glutamate than when growing on glucose, even though the amino acid composition was similar among treatments. If grown on complex substrates, their enzymatic degradation would likely enhance the release of DOM by HP (Aparicio et al., 2015). Further work is needed to explore which would be the main nutritional drivers, e.g. going beyond glucose amendments to testing the effect of the DOM source, of DOM production by HP in the sea.

## Quantity and quality of bacterial DOM

Even though the quantity of DOM released by our two study strains was similar, we could evidence clear differences in its fluorescence signature, which suggests different chemical composition. The production of protein-like FDOM observed for *P. angustum* in the present study, has been previously reported for *Alteromonas macleodii* when grown on glucose (Goto et al., 2017) and for mixed bacterial communities grown on phytoplankton exudates (Romera-Castillo et al., 2011) or on lake water (Cammack et al., 2004). By contrast, the low protein-like FDOM signature in *S. alaskensis* DOM illustrates the importance of bacterial taxonomy for bacterial FDOM quality. *P. angustum* (our study) and *A. macleodii* (Goto et al., 2017) are both fast-growing copiotrophic bacteria, suggesting a link between aspects of bacterial metabolism and protein-like FDOM release. This is in line with the fact that copiotrophs have a higher number of secreted proteins than oligotrophs (Lauro et al., 2009), thus the quantity and composition of the secreted DOM would be likely dependent on the expression and production of these proteins. The production of humic-like DOM by bacteria has been shown using single strains (Shimotori et al., 2012) and experimental incubations with mixed communities (Rochelle-Newall and Fisher, 2002; Romera-Castillo et al., 2011).
275 Humic-like FDOM (peaks A, C and M) is released to the ambient seawater as a
276 byproduct of organic matter remineralization by HP, and has been used as a tracer for
277 refractory DOM in the ocean (Yamashita and Tanoue, 2008; Catalá et al., 2015).

278 Therefore, we hypothesized that DOM produced by *S. alaskensis*, more rich in humic-
279 like compounds, would be less available for HP degradation in our experiments.

280 Few previous published studies have tested bacterial-DOM degradation by in situ HP
281 communities. Both Kramer and Herndl (2004) and Zhang et al. (2015), these latter
282 authors only using the exopolysaccharide fraction of bacterial DOM, concluded that
283 DOM derived from mixed communities was mostly refractory, since no significant
284 growth was detected during 2 to 14 days. However, Lønborg et al. (2009) followed
285 DOC changes over longer time periods (30 d), and concluded that at least a fraction of
286 HP-derived DOM can be utilized by other prokaryotic communities, indicating that HP-
287 DOM is semilabile. This is in accordance with our results, showing that 51% of organic
288 carbon produced by the strains was further used by in situ HP communities (Table 3).

289 **Influence of HP community composition for the degradation of bacterial DOM**

290 In contrast to our hypothesis, differences in the lability of *P. angustum* vs. *S. alaskensis*
291 DOM could not be demonstrated by our biodegradation experiments, as inverse growth
292 patterns on the two strain-derived DOM treatments were observed between coastal
293 communities from April and December (Fig. 2). This suggests that not only the quality
294 of the bacterial DOM, but also the in situ HP community composition would affect
295 bacterial DOM degradation, as previously shown (Nelson and Wear, 2014). Our study
296 strains released DOM that could not be used by themselves, since little bacterial growth
297 of the residual cells was observed even when reinoculating with inorganic nutrients
298 (control treatments in biodegradation experiments, Fig. 2), although this could be due to
299 the possible damage of those cells after freezing. Whether the bacterial DOM can be
300 degraded by single strains, as shown for labile in situ DOM and *Alteromonas macleodii*
301 (Pedler et al., 2014), or whether the complementary metabolic strategies of a diverse
302 community are required, remains to be investigated. The remarkable growth of in situ
303 HP communities was not only due to the glucose that remained in the pdom and sdom
304 treatments, since HP abundance was in both cases higher than in the +glucose
305 treatment. A possibility that we cannot exclude is that this remaining glucose boosted
306 the utilization of other substrates. Differences in HP growth among experiments can be
307 explained not only by differences in the in situ communities but also by in situ DOM
308 variability: In late spring in the Mediterranean Sea, DOM is usually enriched in labile
309 compounds after the spring phytoplankton bloom (Jones et. al., 2013, Ortega-Retuerta
310 et. al., 2018), so the in situ DOM was likely more labile than bacterial derived DOM, in
311 contrast to December, when DOM can have other signatures (i.e. terrestrial-like,
312 Sanchez-Pérez et. al., 2020).

313 In fact, despite non-significant differences in HP growth, consistent patterns of
314 exoenzyme activity were detected: *P. angustum* DOM always promoted high leucine-
315 aminopeptidase activity rates (Fig. 4), in line with its relative enrichment in protein-like
316 FDOM compounds (Table 2). By contrast, *S. alaskensis* DOM consistently promoted
317 relatively high levels of alkaline phosphatase (Fig. 4). This last result was intriguing to
318 us, since inorganic phosphorus was added at the beginning of each biodegradation
319 experiments and thus HP communities were not P-limited. Although high alkaline
320 phosphatase activities have been traditionally interpreted as P limitation (Cotner et al.,
321 1997; Thingstad et al., 1998), HP can use these exoenzymes to release available
322 dissolved organic carbon, as pointed out by previous studies (Benitez-Nelson and
323 Buesseler, 1999; Van Wambeke et al., 2002). Our results suggest that *S. alaskensis*
324 DOM could be rich in phosphorus-containing groups. We could see (only in the coastal
December experiment) that the *S. alaskensis* DOM treatment was the one with a higher proportion of Rhodospirilales. This order has been found in P-limited ocean areas linked to the presence of the carbon-phosphorus lyase pathway for phosphonate degradation (Sosa et al., 2019), in accordance with our hypothesis of P-rich DOM produced by *S. alaskensis*.

Bacterial DOM sustained a lower HP diversity as compared to the in situ DOM (Table 4). In line with this, Noriega-Ortega et al. (2019) compared the solid- phase extractable DOM produced by single bacterial strains with that of marine DOM from the North Pacific and observed higher diversity of compounds in marine DOM respect to bacterial-derived DOM. HP strains were shown to produce thousands of DOM molecules when using a single carbon substrate, however, the DOM chemical diversity produced by a single bacterium with respect to DOM from coastal seawater is likely to be lower. In any case, the significant differences in community composition between treatments suggest population-specific responses to the differences in quality of the DOM produced by *P. angustum* and *S. alaskensis*.

## Bacterial DOM, PGE and carbon fluxes

The growth on strain-derived DOM induced significantly lower prokaryotic growth efficiency (PGE). To the best of our knowledge, our study provides the first measurements of PGE on bacterial-derived DOM, demonstrating that bacterial DOM is preferentially respired. HP do not only promote DOC accumulation in the ocean through the production of refractory DOM (the microbial carbon pump hypothesis shown in previous studies), but would also enhance carbon remineralization (therefore, acting as a CO₂ source). Noteworthy, we did not include DOM production by natural HP communities in our PGE calculations in the biodegradation experiments, and
349 including DOM production in our calculations would lead to even lower PGE values in
350 all treatments.

351 **Changing the paradigm of carbon fluxes through HP in the ocean**

352 DOM release by HP has been repeatedly demonstrated, however, carbon (and other
353 elements) release by HP as DOM is still overlooked when constraining carbon budgets
354 or other elemental fluxes and models in the ocean. For instance, considering that HP
355 produce, on average, 10% of DOC as DOM (average from values in Table 5), in situ HP
356 production estimates (e.g. using isotopes incorporation such as ³H-leucine, (Kirchman et
357 al., 1985) would be underestimated. To better constrain this uncertainty, it is important
358 to know whether DOM release is linearly related to HP biomass production during short
359 incubation times. The release of DOM would also affect estimates of HP growth
360 efficiency. Prokaryotic carbon demand can be calculated as the sum of HP biomass
361 production and respiration or as differences in DOC concentrations in bottle incubations
362 (the dilution approach (delGiorgio and Cole, 1998)), but in both cases PGE would be
363 overestimated by roughly 10%, according to published results. The overestimation
364 could even be higher if, as demonstrated in our study, this 10% of DOM produced by
365 HP would enhance respiration over biomass production. More studies are needed to
366 better understand the dynamics of DOM production by HP and their drivers, and
367 therefore to constrain carbon fluxes through HP.

368

369 **Conclusion and perspectives**

370 In this study, we demonstrate that two marine heterotrophic bacterial strains growing on
371 glucose release remarkable amounts of DOM (compared with biomass production), and
372 that an important fraction of this bacterial derived DOM is available for Mediterranean
373 Sea HP communities. Should this pattern be observed globally, the production of DOM
374 by prokaryotes in the sea would change carbon fluxes through HP towards a decrease in
375 prokaryotic growth efficiency, as well as shaping HP community and exoenzyme
376 activities. While in the present study DOM release is measured in single strains growing
377 on glucose, the uptake of other organic compounds will likely affect the quantity and
378 lability of the released DOM, hence affecting further metabolisms in the ocean. Further
379 work is needed to better constrain the main mechanisms affecting HP DOM production
380 and degradation in the ocean, thus improving our knowledge on carbon sequestration
381 processes such as the microbial carbon pump.

382 **Experimental procedures**

383 **Bacterial strains growth**

384 In order to get dissolved organic matter (DOM) produced by marine bacteria, we grew
385 axenically two strains with contrasting ecology and different taxonomical classes: The
386 first one, *Photobacterium angustum* S14, is a Gammaproteobacterium (Vibrionales,
387 Vibrionaceae) which was isolated from coastal waters (Humphrey et al., 1983). The
388 second strain, *Sphingopyxis alaskensis* RB2256 (Sphingomonoadales,
389 Sphingomonadaceae), is an Alphaproteobacterium isolated from open waters in the
390 North Pacific Ocean (Eguchi et al., 2001). These bacterial strains can be considered as
391 models of copiotrophic (*P. angustum*) and oligotrophic (*S. alaskensis*) life styles (Lauro
392 et al., 2009). The strains were selected based on their capacity to grow on glucose and
393 the laboratory's experience in culturing them (Matallana-Surget et al., 2009; Koedooder
394 et al., 2018). Both strains were preserved in glycerol at -80°C. The stocks were thawed
395 and grown in minimum media, modified from Fegatella et al. (1998), composed of
396 artificial seawater (salinity 35) with glucose as the sole carbon source (≈200 µM C),
397 nitrogen (50:50 mixture of NH₄Cl and NO₃) and phosphorus (NaHPO₄) at a C:N:P ratio
of 45:9:1, equivalent to the internal stoichiometry of bacterial cells (Goldman et al., 1987), and trace amounts of metals and vitamins. In order to minimize the presence of glycerol in the media and get the cells adapted to the growth conditions, two acclimation steps were made, where cells were diluted on fresh media (initial concentration ≈10⁴ cell mL⁻¹) and grown to 1-2 x10⁶ cell mL⁻¹. Purity of the cultures was controlled at each step by plating samples on Marine Agar. Experimental incubations were set up in triplicate in 500 mL flasks, with two contamination controls, consisting of flasks containing culture media but without bacterial inoculation. Incubations were performed at 25°C with rotary agitation (90 rpm) until the stationary phase was reached (controlled several times per day by flow cytometry). No bacterial growth was detected in the contamination controls. Once the stationary phase was reached, the DOM was harvested by gentle filtration through pre-rinsed (Milli-Q water, 250 mL) 0.2 µm polycarbonate filters using an all-glass filtration system. All glass material used for the incubations and filtrations was previously combusted (450°C, 5h). Previous tests (dissolved organic carbon concentration and DOM fluorescence measurements on Milli-Q water filtered using the same device and filters) showed insignificant DOM leaching from the polycarbonate filters (data not shown). In the DOM filtrates, samples for glucose, dissolved organic carbon (DOC), inorganic nutrients, DOM fluorescence and flow cytometry (to check for the presence of cells passing through the filters) were taken, and the remaining DOM was kept at -20°C before being used as substrates for biodegradation experiments (see below). We chose to sample in the early stationary phase to ensure that minimal amounts of glucose were remaining in the DOM extracts for the biodegradation experiments, although at this stage some of the bacterial DOM could derive from cell lysis. Strain cultures p1 and s1 (Fig. 1) were used for the biodegradation experiments 1 and 2, while strain cultures p2
and s2 were used for biodegradation experiment 3. Being aware that freezing could alter some DOM properties, we chose this preservation method due to the inability to perform the biodegradation experiments shortly after the strain incubations were finished.

## 427 Biodegradation experiments

To study how DOM derived from the bacterial strains is degraded by natural marine heterotrophic prokaryotic (HP) communities, re-growth cultures were performed (Table 1). These consisted of 10% dilutions of natural HP communities from different NW Mediterranean Sea locations (water prefiltered by 0.8 µm to remove most phytoplankton and grazers) with 90% DOM (0.2 µm filtrates) from the following sources: DOM sampled from the same waters as the HP communities (in situ surface or deep DOM, ndom treatments) or DOM harvested from the strain cultures explained above: *Photobacterium angustum* (*P. angustum* DOM treatments, pdom) or *Sphingopyxis alaskensis* (*S. alaskensis* DOM treatments, sdom). DOM for ndom treatments was prepared by filtering seawater from the same location as the prokaryotic innocula through 0.2 µm filters as for the bacterial DOM. Inorganic nutrients (nitrogen (50:50 mixture of NH₄Cl and NO₃) and phosphorus (NaHPO₄), final concentrations in supplementary Table 1) were added to each treatment to force carbon limitation, and all treatments were set up in triplicate in 250 mL flasks. Incubations were performed at 18°C in the dark for 7-8 days.

Using this approach, we performed three different experiments: Open sea-March experiment: We selected waters from a station located on the edge of the continental shelf (hereafter termed as “open sea”) in the North Western Mediterranean Sea (42°27′205 N – 03°32′565 E, 600m depth) at two different depths: 5m (surface communities) and 500m (deep communities), in March 2018. These natural HP
communities were grown on in situ DOM (surface or deep DOM) or on *P. angustum*
DOM, with the purpose to evaluate the differences in HP growth depending on the
initial community (surface vs. deep marine communities). For this experiment, samples
for DOC, FDOM and nutrients were taken at the onset and end of the incubations, while
HP abundance was monitored daily by flow cytometry. For the Coastal sea-April
experiment, we collected surface waters from a coastal station (42°29'3N, 3°8'7E, 20m
bottom depth) in April 2018 and grew HP communities on in situ DOM, DOM
produced by *P. angustum*, and DOM produced by *S. alaskensis*, to evaluate growth of
the same HP community on different DOM sources. In this experiment, we monitored
the same variables as in Open sea-April experiment and, in addition, we measured the
exoenzyme activity at the onset, day 3, and end of the experiment. For the Coastal sea-
December experiment, we collected surface waters from the same coastal station as for
biodegradation Experiment 2 (42°29'3N, 3°8'7E, 20m depth), but in December 2018.
The treatments were the same as in May 2018 and thus allowed us to investigate
seasonal differences. We included an additional “+Gluc” treatment to evaluate the effect
of residual glucose amounts in the bacterial DOM sources. In this last experiment, we
monitored the same variables as in biodegradation experiment 2 (DOM, nutrients, HP
abundance and exoenzyme activities) and, additionally, we measured the HP
community structure (16s rRNA Illumina sequencing) at the onset and end of the
experiment.

468 Chemical and biological analyses

469 *Dissolved organic carbon (DOC)*

Samples (10 mL) for DOC were transferred into precombusted glass tubes, acidified
with 85% H₃PO₄ (final pH 2), closed with Teflon lined screw caps and stored in the
472 dark at room temperature until analysis (less than 2 months after sampling). Calibration
473 curves were made using an acetanilide solution ($C_8H_9NO$; M= 135.17 g mol$^{-1}$). DOC
474 was analysed using the high temperature catalytic oxidation (HTCO) technique (Benner
475 and Strom, 1993) with a Shimadzu TOC-V-CSH analyser. Standards of 44–45 µmol C
476 L$^{-1}$, provided by D.A. Hansell and Wenhao Chen (Univ. of Miami), were used to assess
477 the accuracy of the measurements.

478 Glucose

479 Glucose was quantified using a modification of the colorimetric protocol described by
480 Myklestad et al. (1997). Samples (200 µL) were stored in pre-combusted glass vials at -
481 20°C until analysis. All reagents were downscaled to 200 µL samples, and incubations
482 after reagent A additions were performed during 20 min in a 100°C water bath. The
483 reagents were calibrated daily using a standard curve made of D-glucose in the same
484 batch of artificial seawater used for the strain incubations. Milli-Q water and artificial
485 sea water blanks were run daily in triplicate. Glucose concentrations in µmol were
486 converted in carbon equivalent (C-Glucose).

487 Fluorescent dissolved organic matter (FDOM)

488 FDOM samples were freshly (within 1-3 h) analyzed with a Perkin Elmer luminescence
489 spectrometer LS 55 equipped with a xenon discharge lamp, equivalent to 20 kW. Slit
490 widths were 10.0 nm for the wavelengths of excitation and emission. We characterized
491 4 groups of fluorophores using the following excitation/emission (ex/em) pairs (Coble,
492 1996): 280 nm/350 nm (peak T, surrogate of protein-like substances); 250 nm/435 nm
493 (peak A); 340 nm/440 nm (peak C) and 320 nm/410 nm (peak M) the later three due to
494 humic substances of different origins (Coble, 1996). Fluorescence intensities of the
495 peaks were reported in Raman units (R.U.) obtained by dividing the fluorescence units
496 by the Milli-Q blank peak area (Raman scatter) excited at 350 nm.

497 *Dissolved inorganic nutrients*

498 Samples for inorganic nutrient analyses were kept frozen at -20°C. NO₃⁺+NO₂⁻, and PO₄³⁻ were quantified with a segmented flow analyser (Bran Luebbe) with colorimetric detection using methods described in Holmes (1999). The accuracy of the methods was assessed using reference material (Certipur, Merck). The precisions were in the range of 1–4%, and the detection limits were 0.02 µM for NO₃⁺+NO₂⁻, and 0.03 µM for PO₄³⁻. NH₄ samples were collected into pre-rinsed 20 mL HDPE bottles and stored frozen at -20°C before being analysed by fluorometric detection (Holmes, 1999).

505 *Bacterial and heterotrophic prokaryotic abundance*

506 Cell abundances in all experiments were monitored by flow cytometry. Subsamples 507 were fixed with glutaraldehyde (0.5% final concentration) and preserved at -80°C prior 508 to analysis (within days). Cell abundance was measured by using Beckman CytoFLEX 509 and BD FACSCanto flow cytometers equipped with lasers exciting at 488 nm. Samples 510 were thawed and stained with SYBR Green (0.025% (v/v) final concentration). 511 Fluorescent beads (1 µm) were added as internal standards. Samples were run at low 512 speed (10 to 16 µL min⁻¹) for 60 to 90 seconds. Prokaryote communities were identified 513 in plots of side scattered light vs. green fluorescence (del Giorgio et al., 1996).

514 *Extracellular enzyme activities*

515 Extracellular enzyme activities were quantified with the use of fluorogenic substrates 516 (Hoppe, 1983) with the modifications for plate readers described in Sala et al. (2016). 517 Each sample was pipetted in triplicate into 96-well black plates with the following 518 substrates: 4-methylumbelliferyl β-D-glucopyranoside (for β-glucosidase), 4-
519 methylumbelliferyl α-D-glucopyranoside (for α-glucosidase), 4-methylumbelliferyl phosphate (for alkaline phosphatase), and L-leucine-7-amido-4-methyl coumarin (for leu-aminopeptidase) added (final concentration 120 µmol L⁻¹). Fluorescence was measured immediately after addition of the substrate and after incubations (up to 5h with one middle timepoint) in the dark at the same temperature as for the experimental incubations. Fluorescence readings were done with a Victor3 Perkin Elmer spectrofluorometer at 355/450 nm ex/em wavelengths. The increase of fluorescence units during the period of incubation was converted into enzymatic activity with a standard curve prepared with the end products of the reactions, 7-amido-4-methylcoumarin (MCA) for leu-aminopeptidase and 4-methylumbelliferone (MUF) for the rest of enzymes.

530 DNA extraction and sequencing

531 Samples (110 mL) for bacterial diversity were filtered onto 0.2 µm pore size polycarbonate filters. The 0.2 µm filters were stored at -80 °C until analysis. For analysis, frozen filters were cut with sterilized scissors into small strips and vortexed briefly in 750 µL of lysis buffer (50 mmol L⁻¹ Tris hydrochloride pH 8.3, 40 mmol L⁻¹ EDTA and 0.75 mol L⁻¹ sucrose) and then subjected to three freeze-thaw cycles (immersed successively into liquid nitrogen and hot water). Cell lysis was accomplished by an initial incubation for 45 min at 37 °C after adding 41 µL of freshly prepared lysozyme solution (36 mg mL⁻¹), and a second incubation at 55 °C for 1 h after adding 80 µL of 10% sodium dodecyl sulfate and 8 µL of proteinase K (20 mg mL⁻¹). After denaturation and degradation of proteins, DNA was purified using a Quick-DNA™ fungal/bacteria miniprep kit (Zymo Research, Catalog No. D6005) according to the manufacturer's instructions.
The V4-V5 region of the 16S rRNA gene from both fractions was amplified with the primer sets 515F-Y (5'-GTGYCAGCMGCCGCGGTAA) and 926-R (5'-CCGYCAATTYMTTTTRAGTTT) as described in Parada et al. (2016) with a modification to the PCR amplification step. Triplicate 20µL reaction mixtures contained 2 µg DNA, 5 µl KAPA2G Fast HotStart ReadyMix, 0.2 µM forward primer and 0.2 µM reverse primer. Cycling reaction started with a 3 min heating step at 95°C followed by 30 cycles of 95°C for 45 s, 50°C for 45 s, 68°C for 90 s, and a final extension of 68°C for 5 min. The presence of amplification products was confirmed by 1% agarose electrophoresis and triplicate reactions were pooled. The pooled PCR amplicons were purified using Sephadex G-50 Superfine resin (GE Healthcare Bio-Sciences, New Jersey, USA) following the protocol. The purification step aims to desalt the samples and eliminate unincorporated nucleotides and excess PCR primers. 16S rRNA gene amplicons were sequenced with Illumina MiSeq 2 × 250 bp chemistry on one flow-cell at GeT-PlaGe platform (Toulouse, France). A mock community DNA (LGC standards, UK) was used as a standard for subsequent analyses and considered as a DNA sample for all treatments.

All samples from the sequencing run were demultiplexed by GeT-PlaGe and barcodes were trimmed off. Processing of sequences was performed using the DADA2 pipeline (version 1.10) (Callahan et al., 2016) in R (version 3.4.2) with following parameters: `trimLeft=c(19, 20)`, `truncLen=c(240, 200)`, `maxN=0`, `maxEE=c(2, 2)`, `truncQ=2`. Briefly, the pipeline combines the following steps: filtering and trimming, dereplication, sample inference, chimera identification, and merging of paired-end reads. It provides exact amplicon sequence variants (ASVs) from sequencing data with one nucleotide difference instead of building operational taxonomic units (OTUs) based on sequence similarity. ASVs were assigned against SILVA release 132 database (Quast et al.,
568 2013). Singletons and sequences assigned to chloroplast and mitochondria were
569 removed prior to subsequent analyses. Sequences were submitted to NCBI under the
570 accession number PRJNA630555

571

572 **Data analyses**

573 In the strain incubations, we calculated the % of initial DOC transformed to bacterial-
574 derived DOM (%DOC) as the difference between final carbon as glucose (C-Glucose)
575 and dissolved organic carbon (DOC) along the incubations as follows:

$$
576 \quad \%\text{DOC} = \frac{\text{DOC}_{\text{tf}}(\mu\text{mol L}^{-1}) - \text{C-Glucose}_{\text{tf}}(\mu\text{mol L}^{-1})}{\text{DOC}_{t0}(\mu\text{mol L}^{-1})} \quad (1)
$$

577 This equation would be applicable only if at t0, DOC and glucose values were equal.
578 However, there were slight differences between these two pools at t0 in the S.
579 *alaskensis* cultures s1 and s2 (Fig. 1). Because glucose was the sole organic carbon
580 source added to the cultures (final concentration of 200 µM C-glucose), the difference is
581 likely due to the introduction of DOC contamination while manipulating the cultures, or
582 due to analytical differences between the glucose and DOC analyses.

583 Thus, we decided to consider this initial DOC excess (DOC(t0)-C-glucose(t0)) as a non-
584 reactive pool and subtract it from the concentrations at the beginning and end of each
585 incubation. This is the most conservative way to calculate the %DOC released as DOM
586 by the strains, since, alternatively, the strains could have taken up some of this extra
587 DOC and then released a fraction of it back as bacterial DOM.

588 Prokaryotic growth efficiency (PGE) in the biodegradation experiments was calculated
589 as the increase in cell biomass between t0 and tf divided by the decrease in DOC
590 between t0 and tf in the incubations using this formula:

$$
591 \quad \text{PGE (\%)} = \frac{\Delta\text{Biomass (}\mu\text{mol L}^{-1}\text{)}}{\Delta\text{DOC (}\mu\text{mol L}^{-1}\text{)}} \times 100 \quad (2)
$$

592 using a general cell to biomass conversion factor of 20 fg cell⁻¹ (Watson et al., 1977).
593 Differences in HP abundance between treatments in the biodegradation experiments

594 were tested using repeated measures ANOVA and post-hoc pairwise t-tests (package

595 “rstatix” in R). Differences in prokaryotic community structure were tested by

596 ANOSIM tests (package “vegan” in R).

597 Acknowledgements

598 This work was supported by the Caramba project (European commission, H2020-

599 MSCA-IF-2015-703991). Flow cytometric analyses were performed at the BioPic

600 cytometry and imaging platform (Sorbonne University/CNRS). DNA extractions and

601 PCR amplifications were conducted at the Bio2Mar platform (Sorbonne

602 University/CNRS). Dr. Yan Liu kindly provided assistance in PCR and DNA cleaning

603 steps as well as in sequence processing. The 16s samples were sequenced in the INRAE

604 platform. We thank the crew of R/V ‘Nereis II’ and the technicians of the Banyuls

605 observation service for their assistance in getting Mediterranean Sea samples for the

606 biodegradation experiments.

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773 Watson, S.W., Novitsky, T.J., Quinby, H.L., and Valois, F.W. (1977) Determination of bacterial number and biomass in the marine environment. *Appl Envir Microb* **33**: 940-946.

776 Yamashita, Y., and Tanoue, E. (2008) Production of bio-refractory fluorescent dissolved organic matter in the ocean interior. *Nature Geoscience* **1**: 579-582.

778 Zhang, Z., Chen, Y., Wang, R., Cai, R., Fu, Y., and Jiao, N. (2015) The Fate of Marine Bacterial Exopolysaccharide in Natural Marine Microbial Communities. *PLoS One* **10**: e0142690.

781

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**Table 1. Overview of the biodegradation experiments:** Incubation date, treatment abbreviations, dissolved organic matter (DOM) and heterotrophic prokaryote (HP) innocula sources. deep.n: Deep DOM-Deep HP. deep.p: *P. angustum* DOM-Deep HP surf.n: Surface DOM-Surface HP. surf.p: *P. angustum* DOM-Surface HP. ndom: Surface DOM-Surface HP. +gluc: Surface DOM+ glucose-Surface HP. pdom: *P. angustum* DOM-Surface HP. sdom: *S. alaskensis* DOM-Surface HP

789

<table><thead><tr><th>Experiment</th><th>Date</th><th>Treatment</th><th>DOM Source (90% vol)</th><th>HP Innocula</th><th>Abundance DOC Nuts. FDOM</th><th>Enzymes</th><th>16s</th></tr></thead><tbody><tr><td rowspan="4">Open Sea- March</td><td rowspan="4">29 Mar 2018</td><td>deep.n</td><td>500m</td><td>500m</td><td rowspan="4">√</td><td rowspan="4">√</td><td rowspan="4"></td></tr><tr><td>deep.p</td><td><i>P. angustum</i></td><td>“</td></tr><tr><td>surf.n</td><td>5m</td><td>5m</td></tr><tr><td>surf.p</td><td><i>P. angustum</i></td><td>“</td></tr><tr><td rowspan="3">Coastal Sea- April</td><td rowspan="3">10 Apr 2018</td><td>ndom</td><td>5m</td><td>5m</td><td rowspan="3">√</td><td rowspan="3">√</td><td rowspan="3"></td></tr><tr><td>pdom</td><td><i>P. angustum</i></td><td>“</td></tr><tr><td>sdom</td><td><i>S. alaskensis</i></td><td>“</td></tr><tr><td rowspan="4">Coastal Sea- December</td><td rowspan="4">12 Dec 2018</td><td>ndom</td><td>5m</td><td>5m</td><td rowspan="4">√</td><td rowspan="4">√</td><td rowspan="4">√</td></tr><tr><td>+gluc</td><td>5m</td><td>“</td></tr><tr><td></td><td>(+glucose)</td><td></td></tr><tr><td>pdom</td><td><i>P. angustum</i></td><td>“</td></tr><tr><td></td><td>sdom</td><td><i>S. alaskensis</i></td><td>“</td><td></td><td></td><td></td><td></td></tr></tbody></table>

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**Table 2:** Characteristics of bacterial-derived DOM (mean ± standard deviation):
percentage of initial DOC released as bacterial dissolved organic matter (%DOC), final-initial (Δ) values of fluorescent DOM peaks C, M, A (all surrogates for humic-like DOM, x10⁻³ Raman Units) and T (surrogate for protein-like DOM, x10⁻³ Raman Units) and the ratio between Peaks C and T (C/T, indicative of the prevalence of protein-like over humic-like DOM). p: *Photobacterium angustum* s: *Sphingopyxis alaskensis*. p 1 and p 2, and s 1 and s 2 refer to independent experiments (See Experimental Procedures for details).

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<table><thead><tr><th>Experiment</th><th>p 1</th><th>p 2</th><th>s 1</th><th>s 2</th></tr></thead><tbody><tr><td>Date</td><td>03/2018</td><td>07/2018</td><td>03/2018</td><td>11/2018</td></tr><tr><td>%DOC</td><td>22.2±10.6</td><td>10.3±2.1</td><td>6.3±7.1</td><td>10.3±4.6</td></tr><tr><td>Δ FDOM.C</td><td>3.21±0.14</td><td>2.47±0.12</td><td>2.64±1.68</td><td>5.72±0.19</td></tr><tr><td>Δ FDOM.M</td><td>4.84±0.44</td><td>2.79±0.18</td><td>9.03±3.38</td><td>12.88±0.02</td></tr><tr><td>Δ FDOM.A</td><td>4.29±0.55</td><td>5.43±0.41</td><td>13.74±6.63</td><td>16.36±0.52</td></tr><tr><td>Δ FDOM.T</td><td>35.37±1.60</td><td>33.29±1.4</td><td>6.38±0.54</td><td>11.37±0.70</td></tr><tr><td>C/T</td><td>0.09±0.008</td><td>0.07±0.006</td><td>0.42±0.30</td><td>0.50±0.06</td></tr></tbody></table>

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802 Table 3. See horizontal Tables document

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<caption>Table 4. Diversity indexes: Richness, Evenness and Shannon for every treatment of the Biodegradation experiment 3</caption>

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<table><thead><tr><th rowspan="2">Treatment</th><th colspan="2">Richness</th><th colspan="2">Evenness</th><th colspan="2">Shannon</th></tr><tr><th>mean</th><th>SD</th><th>mean</th><th>SD</th><th>mean</th><th>SD</th></tr></thead><tbody><tr><td>Initial</td><td>308</td><td></td><td>0.86</td><td></td><td>4.93</td><td></td></tr><tr><td>Surf HP- Surf DOM</td><td>112</td><td>6</td><td>0.50</td><td>0.02</td><td>2.36</td><td>0.10</td></tr><tr><td>Surf HP- Surf DOM+Gluc</td><td>87</td><td>9</td><td>0.44</td><td>0.04</td><td>1.98</td><td>0.16</td></tr><tr><td>Surf HP- <i>P. angustum</i> DOM</td><td>81</td><td>10</td><td>0.59</td><td>0.03</td><td>2.60</td><td>0.16</td></tr><tr><td>Surf HP- <i>S. alaskensis</i> DOM</td><td>40</td><td>4</td><td>0.46</td><td>0.11</td><td>1.69</td><td>0.39</td></tr></tbody></table>

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813 Table 5: see horizontal Tables document

814
815 **Figure Captions**

816

817 **Figure 1.** Dissolved organic carbon (DOC, µmol L⁻¹, dark grey columns) and glucose-C (µmol C L⁻¹, light grey columns) at the beginning (t0) and end (tf) of the growth experiments of each strain. Whiskers: Standard deviation; p1, p2: *P. angustum* s1, s2: *S. alaskensis* as described on Table 2.

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<table><caption>Table 3. Initial and final values of dissolved organic carbon (DOC) concentration (µmol L⁻¹), ΔDOC per time (µmol L⁻¹ h⁻¹), ΔDOC/DOCT0 (%), ΔBiomass/DOC (%) and DOM fluorescence (protein-like peak T, humic-like peaks A, C and M, Raman Units) in all biodegradation experiments; tf: mean ± SD of biological triplicates</caption><thead><tr><th rowspan="2">Experiment-Treatment</th><th colspan="2">DOC</th><th rowspan="2">ΔDOC/t</th><th rowspan="2">ΔBiomass/DOC (%)</th><th rowspan="2">ΔDOC/DOCT0 (%)</th><th colspan="2">FDOM.C</th><th colspan="2">FDOM.M</th><th colspan="2">FDOM.A</th><th colspan="2">FDOM.T</th></tr><tr><th>t0</th><th>tf</th><th>t0</th><th>tf</th><th>t0</th><th>tf</th><th>t0</th><th>tf</th><th>t0</th><th>tf</th></tr></thead><tbody><tr><td colspan="14"><em>Open Sea-March</em></td></tr><tr><td>Deep HP- Deep DOM</td><td>60.8</td><td>56.2 ± 2.1</td><td>0.04</td><td>8.4 ± 0.5</td><td>7.8</td><td>1.8</td><td>5.0</td><td>7.7</td><td>6.1</td><td>18.9</td><td>14.0</td><td>10.9</td><td>7.2</td></tr><tr><td>Deep HP- <em>P. angustum</em> DOM</td><td>52.7</td><td>30.5 ± 4.5</td><td>0.18</td><td>4.9 ± 1.6</td><td>41.1</td><td>1.6</td><td>5.4</td><td>8.1</td><td>5.6</td><td>14.4</td><td>12.7</td><td>38.7</td><td>28.1</td></tr><tr><td>Surf HP- Surf DOM</td><td>70.9</td><td>61.8 ± 2.8</td><td>0.08</td><td>4.0 ± 1.3</td><td>12.6</td><td>12.9</td><td>7.2</td><td>15.8</td><td>7.5</td><td>83.3</td><td>19.8</td><td>18.8</td><td>8.6</td></tr><tr><td>Surf HP- <em>P. angustum</em> DOM</td><td>72.7</td><td>32.8 ± 8.1</td><td>0.33</td><td>1.4 ± 0.3</td><td>54.6</td><td>2.2</td><td>5.7</td><td>7.9</td><td>5.5</td><td>15.3</td><td>12.8</td><td>34.1</td><td>22.0</td></tr><tr><td colspan="14"><em>Coastal Sea-April</em></td></tr><tr><td>Surf HP- Surf DOM</td><td>99.2</td><td>75.5 ± 12.8</td><td>0.16</td><td>3.3 ± 1.1</td><td>23.4</td><td>10.3</td><td>16.2</td><td>11.6</td><td>12.1</td><td>27.4</td><td>29.1</td><td>11.3</td><td>13.2</td></tr><tr><td>Surf HP- <em>P. angustum</em> DOM</td><td>98.1</td><td>34.1 ± 15.5</td><td>0.45</td><td>2.0 ± 0.2</td><td>65.3</td><td>9.2</td><td>11.8</td><td>16.5</td><td>6.8</td><td>25.1</td><td>15.2</td><td>53.7</td><td>14.0</td></tr><tr><td>Surf HP- <em>S. alaskensis</em> DOM</td><td>76.0</td><td>57.0 ± 4.1</td><td>0.13</td><td>2.5 ± 0.7</td><td>25.0</td><td>16.5</td><td>21.9</td><td>32.3</td><td>17.1</td><td>41.2</td><td>16.9</td><td>16.5</td><td>14.4</td></tr><tr><td colspan="14"><em>Coastal Sea-December</em></td></tr><tr><td>Surf HP- Surf DOM</td><td>83.0</td><td>74.1 ± 2.3</td><td>0.05</td><td>2.0 ± 0.6</td><td>10.8</td><td>11.2</td><td>11.2</td><td>12.1</td><td>12.5</td><td>28.4</td><td>29.2</td><td>14.3</td><td>11.2</td></tr><tr><td>Surf HP- Surf DOM+Gluc</td><td>107.3</td><td>78.3 ± 2.4</td><td>0.18</td><td>1.9 ± 0.2</td><td>27.3</td><td>11.5</td><td>10.7</td><td>11.9</td><td>11.3</td><td>29.1</td><td>27.2</td><td>13.2</td><td>12.2</td></tr><tr><td>Surf HP- <em>P. angustum</em> DOM</td><td>90.1</td><td>38.0 ± 2.4</td><td>0.31</td><td>9.2 ± 3.4</td><td>57.8</td><td>6.5</td><td>6.3</td><td>7.6</td><td>8.4</td><td>15.5</td><td>19.7</td><td>35.3</td><td>37.0</td></tr><tr><td>Surf HP- <em>S. alaskensis</em> DOM</td><td>110.0</td><td>39.1 ± 4.0</td><td>0.43</td><td>6.4 ± 1.7</td><td>64.5</td><td>8.4</td><td>9.0</td><td>15.2</td><td>13.5</td><td>23.4</td><td>22.8</td><td>13.3</td><td>13.5</td></tr></tbody></table>
<table><caption>Table 5. Review of previous estimates of dissolved organic matter (DOM) vs. biomass production by heterotrophic prokaryotes as percentage of the initial dissolved organic carbon (DOC) concentration</caption><thead><tr><th colspan="9">Innocula from mixed bacterial communities</th></tr><tr><th>Study</th><th>Substrate used</th><th>Procedence</th><th>DOC Initial (µmol L<sup>-1</sup>)</th><th>Time (d)</th><th>DOC final (µmol L<sup>-1</sup>)</th><th>Biomass final (µmol L<sup>-1</sup>)</th><th>%DOC Released as DOM</th><th>%DOC into biomass</th></tr></thead><tbody><tr><td>(Ogawa et al., 2001)</td><td>Glucose</td><td>Coastal (Gulf of Mexico)</td><td>208</td><td>2</td><td>30</td><td>na</td><td>15.0</td><td>7.0</td></tr><tr><td>“</td><td>Glutamate</td><td>Coastal (Sagami Bay)</td><td>132</td><td>2</td><td>18</td><td>na</td><td>13.0</td><td>22.0</td></tr><tr><td></td><td>Glucose</td><td>Coastal (North Sea)</td><td>220</td><td>5</td><td>24</td><td>13.3</td><td>10.9</td><td>6.0</td></tr><tr><td>(Kawasaki and Benner, 2006)</td><td>Glucose</td><td>Lake (Murray)</td><td>529</td><td>1</td><td>37</td><td>8</td><td>7.0</td><td>1.5</td></tr><tr><td>“</td><td>Glucose</td><td>Estuary (Winyah Bay)</td><td>513</td><td>2</td><td>115</td><td>47</td><td>22.4</td><td>9.2</td></tr><tr><td>“</td><td>Glucose</td><td>Coastal (Folly Beach)</td><td>310</td><td>2</td><td>25</td><td>0.83</td><td>8.1</td><td>0.3</td></tr><tr><td>(Lønborg et al., 2009)</td><td>Glucose</td><td>Fjord (Loch Creran)</td><td>na</td><td>30</td><td>na</td><td>na</td><td>11</td><td>na</td></tr><tr><td>(Koch et al., 2014)</td><td>Glucose</td><td>Antarctic</td><td>326</td><td>21</td><td>45</td><td>30</td><td>13.8</td><td>9.2</td></tr><tr><td>(Lechtenfeld et al., 2015)</td><td>Glucose+Glutamic Acid</td><td>Coastal seawater</td><td>283</td><td>1.5</td><td>10</td><td>6.7</td><td>3.5</td><td>2.4</td></tr><tr><td>(Arai et al., 2017)</td><td>Glucose</td><td>Coastal (Shizoka, Japan)</td><td>833</td><td>5</td><td>60.4</td><td>272</td><td>7.3</td><td>32.4</td></tr></tbody><thead><tr><th colspan="9">Innocula from single bacterial strains</th></tr><tr><th>Study</th><th>Substrate used</th><th>Strain</th><th>DOC initial (µmol L<sup>-1</sup>)</th><th>Time (d)</th><th>DOC final (µmol L<sup>-1</sup>)</th><th>Biomass final (µmol L<sup>-1</sup>)</th><th>%DOC Released as DOM</th><th>%DOC into biomass</th></tr></thead><tbody><tr><td>(Romano et al., 2014)</td><td>Glucose, P limited</td><td>Pseudovibrio sp. FO-BEG1</td><td>60000</td><td></td><td>na</td><td>2000</td><td>0.9*</td><td>3.3</td></tr><tr><td>“</td><td>Glucose, P replete</td><td>“</td><td></td><td></td><td></td><td></td><td>0.2*</td><td></td></tr><tr><td>(Zhang et al., 2015)</td><td>Glucose</td><td>Alteromonas sp. JL2810</td><td>3333</td><td>5</td><td></td><td></td><td>8**</td><td></td></tr><tr><td>(Goto et al., 2017)</td><td>Glucose</td><td>Alteromonas macleodii</td><td>995</td><td>1</td><td>58</td><td>6.1</td><td>5.8</td><td>0.6</td></tr><tr><td>(Noriega-Ortega et al., 2019)</td><td>Acetate</td><td>Phaeobacter inhibens</td><td>70050</td><td>na</td><td>2400</td><td>na</td><td>3.4</td><td>na</td></tr><tr><td>Present Study</td><td>Glucose</td><td>Photobacterium angustum</td><td>198</td><td></td><td>55</td><td>15.4</td><td>16.3</td><td>4.4</td></tr><tr><td></td><td></td><td>Sphingopyxis alaskensis</td><td>245</td><td>7</td><td>72</td><td>8.4</td><td>8.5</td><td>7.6</td></tr></tbody></table>

bdl: below detection limit; na: not available (not measured or not reported in the paper)

* Only SPE extractable fraction quantified

** Only exopolysaccharide quantified
<figure><img src="image_3.png" /></figure>
<figure><img src="image_4.png" /></figure>
<figure><img src="image_5.png" /><figcaption>**Figure 1.** Dissolved organic carbon (DOC, µmol L⁻¹, dark grey columns) and glucose-C (µmol C L⁻¹, light grey columns) at the beginning (t0) and end (tf) of the growth experiments of each strain. Whiskers: Standard deviation; p1, p2: *P. angustum* s1, s2: *S. alaskensis* as described on Table 2.</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Figure 2.** Changes in prokaryotic heterotrophic abundance (PHA, cells mL⁻¹) over incubation time in the three biodegradation experiments. Open sea-March (A): deep.n: deep water HP and DOM, deep.p: deep water HP, *P. angustum* DOM. surf.n: surface water HP and DOM, surface.p: surface water HP, *P. angustum* DOM. Coastal sea-April (B) and coastal sea-December (C): ndom: surface water HP and DOM; +gluc: surface water HP and DOM, +20 µm L⁻¹ glucose-C added. pdom: surface water HP, *P. angustum* DOM. sdom: surface water HP, *S. alaskensis* DOM. cont.p and cont.s: *P. angustum* and *S. alaskensis* DOM, no HP inoculum added</figcaption></figure>
<figure><img src="image_7.png" /><figcaption>**Figure 3.** Prokaryotic Growth efficiency (PGE, %) in the biodegradation experiments (Biodeg). Open Sea-March: deep.n: deep water HP and in situ DOM, deep.p: deep water HP, *P angustum* DOM. surf.n: surface water HP and in situ DOM, deep.p: surface water HP, *P angustum* DOM. Coastal sea experiments: ndom: surface water HP and in situ DOM; +gluc: surface water HP and DOM, +20 µm L⁻¹ glucose-C added. pdom: surface water HP, *P angustum* DOM. sdom: surface water HP, *S. alaskensis* DOM</figcaption></figure>
<figure><img src="image_8.png" /><figcaption>**Figure 4.** Changes in exoenzymatic activity over time in coastal biodegradation experiments (April: a, b, c, d. December: e, f, g, h). Columns: different study enzymes. Rows: Different experiments</figcaption></figure>
<figure><img src="image_9.png" /><figcaption>**Figure 5.** Left panel: MDS plot of all treatments and replicates based on bray Curtis dissimilarity. Differences in prokaryotic community structure among treatments are significantly (ANOSIM test, p < 0.05). Right panel: Changes in prokaryotic community composition based in 16s rRNA Illumina sequencing in the coastal-December biodegradation experiment. Only orders having ASV's with a relative abundance higher than 2% are plotted.</figcaption></figure>
1 Supplementary Table 1. Inorganic nutrients (phosphate, nitrate+nitrite, ammonium

2 µmol L⁻¹) at the onset (t0) and end (tf) of all strain incubations and biodegradation

3 experiments.

4

<table><thead><tr><th>Strain</th><th>Date</th><th>PO<sub>4</sub> t0</th><th>PO<sub>4</sub> tf</th><th>NO<sub>3</sub> t0</th><th>NO<sub>3</sub> tf</th><th>NH<sub>4</sub> t0</th><th>NH<sub>4</sub> tf</th></tr></thead><tbody><tr><td rowspan="2">S. alaskensis</td><td>mar-18</td><td>2.5</td><td>1.4</td><td>26</td><td>23.4</td><td>25</td><td>7.5</td></tr><tr><td>nov-18</td><td>3.4</td><td>2.5</td><td>28</td><td>26.8</td><td>18.9</td><td>10.5</td></tr><tr><td rowspan="2">P. angustum</td><td>mar-18</td><td>2.8</td><td>1.4</td><td>19.9</td><td>20</td><td>20.3</td><td>7.1</td></tr><tr><td>jul-18</td><td>2.8</td><td>1.4</td><td>23.8</td><td>23.3</td><td>nd</td><td>nd</td></tr><tr><th>Experiment</th><th>Treat</th><th>PO<sub>4</sub> t0</th><th>PO<sub>4</sub> tf</th><th>NO<sub>3</sub> t0</th><th>NO<sub>3</sub> tf</th><th>NH<sub>4</sub> t0</th><th>NH<sub>4</sub> tf</th></tr><tr><td rowspan="4">Open/March</td><td>deep.n</td><td>1.3</td><td>1.23 ± 0.1</td><td>14.3</td><td>13.42 ± 0.8</td><td>nd</td><td>8.94 ± 0.5</td></tr><tr><td>deep.p</td><td>2.0</td><td>1.87 ± 0.3</td><td>27.6</td><td>24.8 ± 2.3</td><td>10.6</td><td>8.47 ± 2.3</td></tr><tr><td>surf.n</td><td>1.3</td><td>1.12 ± 0.1</td><td>13.5</td><td>11.60 ± 0.2</td><td>9.2</td><td>8.94 ± 0.5</td></tr><tr><td>surf.p</td><td>2.5</td><td>1.77 ± 0.4</td><td>28.2</td><td>27.91 ± 4.2</td><td>11.9</td><td>9.57 ± 0.6</td></tr><tr><td rowspan="3">Coastal/ April</td><td>ndom</td><td>1.2</td><td>0.90 ± 0.1</td><td>12.1</td><td>11.33 ± 0.2</td><td>nd</td><td>nd</td></tr><tr><td>pdom</td><td>2.6</td><td>1.63 ± 0.1</td><td>26.7</td><td>25.13 ± 0.5</td><td>nd</td><td>nd</td></tr><tr><td>sdom</td><td>2.0</td><td>1.66 ± 1.9</td><td>31.7</td><td>29.79 ± 0.4</td><td>nd</td><td>nd</td></tr><tr><td rowspan="4">Coastal / Dec</td><td>ndom</td><td>1.6</td><td>1.55 ± 0.1</td><td>9.3</td><td>8.62 ± 0.1</td><td>7.3</td><td>6.86 ± 0.3</td></tr><tr><td>+gluc</td><td>1.4</td><td>1.21 ± 0.0</td><td>9.9</td><td>9.43 ± 0.2</td><td>8.2</td><td>6.45 ± 0.1</td></tr><tr><td>pdom</td><td>2.5</td><td>2.03 ± 0.1</td><td>31.2</td><td>29.64 ± 0.2</td><td>14.6</td><td>12.57 ± 0.4</td></tr><tr><td>sdom</td><td>3.3</td><td>2.54 ± 0.1</td><td>32.6</td><td>30.53 ± 0.5</td><td>16.2</td><td>11.12 ± 0.3</td></tr></tbody></table>

5
Supplementary Figure 1. Changes in cell abundance in the strain growth experiments.

Symbols and whiskers are, respectively, averages and standard deviations of three

biological triplicates. Samples for DOM characterization were taken at the t0 and t final

points in each growth curve. Note the log-10 scale in the y axis

<figure><img src="image_10.png" /></figure>
13 **Supplementary Table 2.** Results of the repeated measures ANOVA tests to compare
14 the changes in cell abundance over the different biodegradation experiments and
15 treatments. DFn: Degrees of freedom in the numerator. DFd: Degrees of freedom in the
denominator. F: F-value p: level of significance. ges: Generalized Eta-Squared measure
16 of effect size.
17

<table><thead><tr><th colspan="6">Open Sea/March</th></tr><tr><th>Effect</th><th>DFn</th><th>DFd</th><th>F</th><th>p</th><th>ges</th></tr></thead><tbody><tr><td>treatment</td><td>3</td><td>6</td><td>25.973</td><td>0.000778</td><td>0.806</td></tr><tr><td>time</td><td>4</td><td>8</td><td>58.986</td><td>5.63E-06</td><td>0.806</td></tr><tr><td>treat:time</td><td>12</td><td>24</td><td>9.901</td><td>1.35E-06</td><td>0.697</td></tr></tbody></table>

<table><thead><tr><th colspan="6">Coastal Sea/April</th></tr><tr><th>Effect</th><th>DFn</th><th>DFd</th><th>F</th><th>p</th><th>ges</th></tr></thead><tbody><tr><td>treatment</td><td>2</td><td>4</td><td>12.73</td><td>0.018</td><td>0.649</td></tr><tr><td>time</td><td>6</td><td>12</td><td>75.287</td><td>8.04E-09</td><td>0.877</td></tr><tr><td>treat:time</td><td>12</td><td>24</td><td>7.134</td><td>2.44E-05</td><td>0.603</td></tr></tbody></table>

<table><thead><tr><th colspan="6">Coastal Sea/December</th></tr><tr><th>Effect</th><th>DFn</th><th>DFd</th><th>F</th><th>p</th><th>ges</th></tr></thead><tbody><tr><td>treatment</td><td>3</td><td>6</td><td>7.242</td><td>0.02</td><td>0.602</td></tr><tr><td>time</td><td>6</td><td>12</td><td>67.137</td><td>1.56E-08</td><td>0.729</td></tr><tr><td>treat:time</td><td>18</td><td>36</td><td>10.226</td><td>2.87E-09</td><td>0.561</td></tr></tbody></table>

18

19
Supplementary Table 3. Results of pairwise t-tests between each treatment and time in the three biodegradation experiments. Different letters denote significantly different treatments at p < 0.05

23

<table><thead><tr><th colspan="5">Open Sea/March</th></tr><tr><th>Time/Treatment</th><th>deep.n</th><th>deep.p</th><th>surf.n</th><th>surf.p</th></tr></thead><tbody><tr><td>1</td><td>a</td><td>a</td><td>b</td><td>c</td></tr><tr><td>2</td><td>a</td><td>a</td><td>ab</td><td>b</td></tr><tr><td>3</td><td>a</td><td>ac</td><td>b</td><td>c</td></tr><tr><td>4</td><td>a</td><td>a</td><td>a</td><td>b</td></tr><tr><td>5</td><td>a</td><td>ab</td><td>ab</td><td>b</td></tr></tbody></table>

<table><thead><tr><th colspan="4">Coastal Sea/April</th></tr><tr><th>Time/Treatment</th><th>ndom</th><th>pdom</th><th>sdom</th></tr></thead><tbody><tr><td>1</td><td>a</td><td>b</td><td>ab</td></tr><tr><td>2</td><td>a</td><td>a</td><td>b</td></tr><tr><td>3</td><td>a</td><td>ab</td><td>b</td></tr><tr><td>4</td><td>a</td><td>ab</td><td>b</td></tr><tr><td>5</td><td>a</td><td>a</td><td>a</td></tr><tr><td>6</td><td>a</td><td>b</td><td>ab</td></tr><tr><td>7</td><td>a</td><td>a</td><td>a</td></tr></tbody></table>

<table><thead><tr><th colspan="5">Coastal Sea/December</th></tr><tr><th>Time/Treatment</th><th>ndom</th><th>+gluc</th><th>pdom</th><th>sdom</th></tr></thead><tbody><tr><td>1</td><td>a</td><td>b</td><td>c</td><td>d</td></tr><tr><td>2</td><td>a</td><td>b</td><td>c</td><td>b</td></tr><tr><td>3</td><td>a</td><td>b</td><td>ab</td><td>ab</td></tr><tr><td>4</td><td>a</td><td>ab</td><td>ab</td><td>b</td></tr><tr><td>5</td><td>a</td><td>b</td><td>abc</td><td>c</td></tr><tr><td>6</td><td>a</td><td>a</td><td>ab</td><td>b</td></tr><tr><td>7</td><td>a</td><td>ab</td><td>bc</td><td>c</td></tr></tbody></table>

24

25
<table><caption>Supplementary Table 4. Composition of the minimum media used for strain growth</caption><thead><tr><th></th><th>Final Concentration (g L<sup>-1</sup>)</th></tr></thead><tbody><tr><td><strong>Sea Salts</strong></td><td></td></tr><tr><td>NaCl</td><td>24</td></tr><tr><td>Na<sub>2</sub>SO<sub>4</sub></td><td>4</td></tr><tr><td>KCl</td><td>0.68</td></tr><tr><td>KBr</td><td>0.1</td></tr><tr><td>H<sub>3</sub>BO<sub>3</sub></td><td>0.025</td></tr><tr><td>NaF</td><td>0.002</td></tr><tr><td>MgCl<sub>2</sub>.6H<sub>2</sub>O</td><td>10.8</td></tr><tr><td>CaCl<sub>2</sub>.2H<sub>2</sub>O</td><td>1.5</td></tr><tr><td>SrCl<sub>2</sub>.6H<sub>2</sub>O</td><td>0.024</td></tr><tr><td>NaHCO<sub>3</sub></td><td>0.2</td></tr><tr><td><strong>Macronutrients</strong></td><td>Final Concentration (µM)</td></tr><tr><td>Glucose</td><td>33 µM (=200 µM C)</td></tr><tr><td>NaNO<sub>3</sub></td><td>22 µM N</td></tr><tr><td>NH<sub>4</sub>Cl</td><td>22 µM N</td></tr><tr><td>NaHPO<sub>4</sub></td><td>4.4 µM P</td></tr><tr><td><strong>Trace Metals</strong></td><td>Final Concentration (µg L<sup>-1</sup>)</td></tr><tr><td>CuCl<sub>2</sub>.2H<sub>2</sub>O</td><td>0.015</td></tr><tr><td>NiCl<sub>2</sub>.H<sub>2</sub>O</td><td>0.025</td></tr><tr><td>Na<sub>2</sub>MOO<sub>4</sub>.2H<sub>2</sub>O</td><td>0.025</td></tr><tr><td>ZnSO<sub>4</sub>.7H<sub>2</sub>O</td><td>0.07</td></tr><tr><td>MnCl<sub>2</sub>.4H<sub>2</sub>O</td><td>0.1</td></tr><tr><td>CoCl<sub>2</sub>.6H<sub>2</sub>O</td><td>0.12</td></tr><tr><td>FeCl<sub>3</sub>.6H<sub>2</sub>O</td><td>4</td></tr><tr><td>EDTA</td><td>2 (0.068 µM C)</td></tr><tr><td><strong>Vitamins</strong></td><td>Final Concentration (µg L<sup>-1</sup>/µM)</td></tr><tr><td>P-aminobenzoic acid</td><td>0.05 / 2.6</td></tr><tr><td>D-biotin</td><td>0.02 / 0.8</td></tr><tr><td>Folic Acid</td><td>0.02 / 0.9</td></tr><tr><td>Niacinamide</td><td>0.02 / 0.9</td></tr><tr><td>D-pantothenic acid</td><td>0.05 / 1.8</td></tr><tr><td>Pyridoxal</td><td>0.02 / 0.8</td></tr><tr><td>Pyridoxamine</td><td>0.10 / 3.3</td></tr><tr><td>Pyridoxine</td><td>0.10 / 3.9</td></tr><tr><td>Riboflavin</td><td>0.05 / 2.3</td></tr><tr><td>Thiamine</td><td>0.02 / 0.9</td></tr><tr><td>D L-6,8-thioctic acid</td><td>0.02 / 0.7</td></tr><tr><td>Vitamin B12</td><td>0.05 / 2.3</td></tr><tr><td>Nicotinamide</td><td>0.05 / 2.5</td></tr></tbody></table>