The strain A. macleodii ATCC 27126 was obtained from the Japan Collection of Microorganisms, RIKEN Bio Resource Center (Tsukuba, Ibaraki, Japan), and was used as a model marine bacterial isolate.

Prior to the present experimental setup, 100 μL frozen stock of A. macleodii was inoculated onto 100 mL artificial seawater-based Aquil medium (Price et al., 1989) supplemented with glucose (1 mmol C L^-1) at 25°C for 24 h. Then, the medium was transferred onto new Aquil medium and incubated under the same conditions as the pre-culture medium. After 24 h of incubation, the culture was used as inoculum for the present experiment.

Alteromonas macleodii was cultivated in modified organic carbon-free Aquil medium (Price et al., 1989) (Table 1). Anhydrous salts used for artificial seawater (NaCl, Na₂SO₄, KCl, KBr) were combusted at 450°C for 4 h. NaH₂PO₄⋅H₂O (10 μmol P L^-1) and NaNO₃ (161 μmol N L^-1) were added as major nutrients. The media for control and experimental treatments were prepared without and with glucose (1 mmol C L^-1) in each triplicate, respectively. The A. macleodii inoculum was added to the medium of each treatment condition at a 1:1000 dilution. Then, each treatment condition with A. macleodii inoculum was dispensed into acid-washed (1 M HCl) 250 mL polyethylene terephthalate bottles. The bottle was filled with 100 mL of culture medium, and thus, gas phase was maintained in the bottles to keep oxygen at a high enough level to catabolize all of the added glucose. Incubations of the experimental and control treatments were conducted in the dark and in an incubator (CN-40, Mitsubishi-Engineering Co.) in which the temperature was maintained at 25°C. To determine the bacterial abundance, DOC concentration and DOM optical property, triplicate bottles were sampled at 0, 6, 12, 18 24, 72, 120, and 168 h in the experimental treatment and at 0 and 168 hours in the control treatment.

Bacterial cell density was measured with an EPICS flow cytometer (XL ADC system, Beckman Coulter) equipped with a 15 mW air-cooled laser exciting at 488 nm, according to the protocol of Tada and Suzuki (2016). Samples were fixed in paraformaldehyde [2% (vol/vol) final concentration] and preserved at –25°C. Just before analysis, samples were stained with SYBR Gold (SYBR Gold Nucleic Acid Gel Stain, Life technologies) at a final concentration of 10^-4 commercial stock solution for at least 15 min. To calculate the flow rate, 2 μm fluorescent beads (Fluoresbrite YG Carboxylate Microspheres 2.00 μm, Polysciences, Inc.) were added to the flow samples. We used a low flow rate mode and analyzed the samples until twenty thousand particles were counted or the measurement time reached 5 minutes. The measured values were corrected by blank value subtraction; the blank was measured with artificial seawater filtered with 0.2-μm pore size cellulose membrane filters (DISMIC-25AS 0.20 μm, ADVANTEC).

Samples for DOM analyses were filtered through pre-combusted (450°C, 3 h) glass fiber filters with a nominal pore size of 0.3 μm (GF75, Whatman) under gentle vacuum (<0.02 MPa) at each incubation time to remove particles, including A. macleodii. The filtrate was collected into a pre-combusted (450°C, 3 h) glass vial with teflon-lined cap and was preserved at –25°C until analysis.

The DOC concentration was determined by high-temperature catalytic oxidation with a total organic carbon analyzer (TOC-V CSH, Shimadzu). The DOC concentrations were calculated using the standard curve of potassium hydrogen phthalate solution, which was determined daily. The accuracy and consistency of the measured DOC concentrations were checked by a deep seawater reference sample (Hansell Laboratory, University of Miami), which was assessed daily.

Excitation–emission matrix was measured using a fluorometer (FluoroMax-4, Horiba) according to the procedure of Tanaka et al. (2014). Samples were allowed to reach near room temperature before the EEM measurements were undertaken. Forty-one emission scans from 290 to 600 nm taken at 2-nm intervals were acquired for the excitation wavelengths between 250 and 450 nm at 5-nm intervals. The bandpass was set to 5 nm for both excitation and emission. The fluorescence spectra were scanned with a 0.25 s integration time and acquired in the S/R mode. Several post-acquisition steps were involved in the correction of the fluorescence spectra, including instrumental bias correction and corrections of inner filter effect using absorbance. Following this, the EEMs of Milli-Q water were subtracted from those of the samples, and fluorescence intensities in EEMs were converted to Raman Units (RU) with the peak areas of Raman scatter at 350 nm excitation (Lawaetz and Stedmon, 2009). RU can be converted to quinine sulfate units (QSU) by using the equation QSU = RU/0.0767 (Lawaetz and Stedmon, 2009). The absorbance spectrum of each sample for correction of inner filter effect was measured with a Shimadzu UV-1800 spectrophotometer in a 1-cm quartz cuvette according to Yamashita et al. (2013). Because nitrate has relatively high absorbance in the UV-B region (Catalá et al., 2016) at high concentrations (e.g., 161 μmol N L^-1 in the medium), investigations of changes in the absorption spectrum associated with A. macleodii incubation were abandoned. The contour of EEMs was plotted by R (version 3.2.3) (R Development Core Team, 2015).

The abundance of A. macleodii increased exponentially during the first 24 h in the experimental treatment (Figure 1A). The cell density was 4.1 × 10⁴ cells mL^-1 at the initiation of incubation (t = 0) and increased up to 3.7 ± 1.8 × 10⁶ cells mL^-1 after 24 h. The cell density then remained stable on a logarithmic scale until the end of the experiment (Figure 1A). Therefore, the periods of 0–24 h and 24–168 h were defined as the exponential growth and stationary phases, respectively. The average specific growth rate was 0.19 ± 0.10 h^-1 during the exponential growth phase. In the control treatment, cell density increased approximately twofold during the incubation (from 3.4 × 10⁴ cells mL^-1 at the initiation of the incubation to 8.0 ± 0.5 × 10⁴ cells mL^-1 at the end of the incubation), indicating that bacterial growth caused by the medium substrate and carbon storage in the cells of the inoculum was much lower than that caused by glucose in the experimental treatment.

The DOC concentration gradually decreased during the early part of the exponential growth phase (first 18-h) from 995 μmol C L^-1 to 830 ± 28 μmol C L^-1 (Figure 1B) and then drastically decreased to 58 ± 15 μmol C L^-1 in the latter part of the exponential growth phase (18–24 h). There was a larger relative decrease in DOC concentration normalized by bacterial cell density between 18 and 24 h (0.29 mmol C cell^-1) than between 0 and 18 h (0.17 mmol C cell^-1), thus suggesting a substantial increase in A. macleodii cell volume between 18 and 24 h. The DOC concentration was almost constant (from 51 ± 4 μmol C L^-1 to 89 ± 29 μmol C L^-1) over the stationary phase and was 51 ± 4 μmol C L^-1 at the end of the experimental incubations (168 h).

The DOC concentrations in the control treatment were 1–2 orders of magnitude lower than those in the experimental treatment and were 7.6 μmol C L^-1 and 9.6 ± 1.5 μmol C L^-1 at the beginning and end of the incubation, respectively. The DOC concentrations detected in the control experiment were possibly derived from the medium and bacterial inoculum.

Fluorescent peaks found in EEMs at the 24-h time point and the end of the experimental treatment are shown in Figures 2B,D, respectively. Because EEMs at the beginning of the experimental treatment (Figure 2A) and at the end of the control treatment (Figure 2E) did not show distinct fluorescent peaks, it can be concluded that fluorescent peaks found for the experimental treatment were derived from A. macleodii and not from the incubation medium or contamination during incubation.

Five fluorescent peaks were defined from EEMs obtained at the end of the experimental treatment (Figure 2D and Table 2). Two peaks were characterized as protein-like fluorescent peaks similar to the aromatic amino acids tyrosine and tryptophan (Coble, 1996; Mayer et al., 1999). From EEMs observed during the experimental treatment (Figures 2B–D), the excitation and emission wavelength (Ex/Em) of tyrosine-like and tryptophan-like peaks were defined to be 275/300 nm and 275/330 nm, respectively. Three peaks could be categorized to humic-like fluorophores. Two of them had emission maximum longer than 500 nm (H1: Ex/Em = 270/520 nm, H3: Ex/Em = 425/520 nm) and have usually been defined to be terrestrial humic-like FDOM in coastal environments (Stedmon and Markager, 2005). The combined fluorescence characteristics of H1 and H3 were similar to that found in the most abundant component in the humic acid fraction extracted from sediments/soils (He et al., 2006; Santín et al., 2009). The other humic-like fluorophore (H2: Ex/Em = 315/400 nm), which had a shorter emission wavelength than H1 and H3, was similar to traditionally defined marine/microbial humic-like FDOM (Coble, 1996, 2007) and was also similar to a fluorophore excreted by cultured phytoplankton (Romera-Castillo et al., 2010).

The fluorescence intensities of two protein-like peaks, namely tyrosine-like and tryptophan-like peaks, increased during the exponential growth phase, particularly during the period of 18–24 h (Figure 3), in which the DOC concentration drastically decreased (Figure 1). Tyrosine-like fluorescence intensity increased during the early part of the stationary phase (24–72 h, Figure 3; Student’s t-test, p < 0.05) and was tended to decrease during the latter part of the stationary phase (72–168 h; Student’s t-test, p > 0.05). Tryptophan-like fluorescence intensity decreased significantly during the latter part of the stationary phase (Figure 3; Student’s t-test, p < 0.05).

Tyrosine-like and tryptophan-like fluorophores exhibited fluorescence peaks with an excitation wavelength of 275 nm (Coble, 2007). It is difficult to quantitatively evaluate changes in protein-like fluorescence intensities, because the emission spectra of tyrosine and tryptophan molecules overlap (Mayer et al., 1999; Lakowicz, 2006). Thus, in this study, the emission spectrum at an excitation wavelength of 275 nm was used to evaluate the dominant protein-like fluorophore during A. macleodii incubation (Figure 4). The emission spectra were almost the same during 0–18 h, but the fluorescence intensities at wavelengths corresponding to protein-like fluorescence (approximately 290–370 nm) were slightly increased with incubation time (Figures 3, 4). A single fluorophore peaked at 330 nm appeared in 24 h, indicating that tryptophan-like fluorophore was mainly produced during the exponential growth phase (Figures 2B, 4). It seemed that two fluorophores peaked at 330 and 300 nm were the main components of the spectrum at 72 h, indicating that both tryptophan-like and tyrosine-like fluorophores were produced during the early part of the stationary phase (Figures 2C, 4). Although fluorescence intensities corresponding to protein-like fluorophores decreased entirely during 72–168 h, the decrease in the tryptophan-like fluorophore (peak at 330 nm) was greater than that of the tyrosine-like fluorophore (peak at 300 nm) (Figures 2D, 4). Furthermore, the tyrosine-like fluorophore became a major peak at 168 h. The fluorescence intensity at 300 nm should be composed of the peak maximum of tyrosine-like fluorophore and the peak edge of tryptophan-like fluorophore. Thus, changes in the shape of emission spectrum observed during 72–168 h imply that the tryptophan-like fluorophore decreased but the tyrosine-like fluorophore was relatively stable during the latter part of the stationary phase.

The fluorescence intensities of all humic-like fluorophores increased during the exponential growth phase (Figure 5). The changes in fluorescence intensity during the stationary phase varied among the humic-like fluorophores (Figure 5). Two humic-like fluorophores with emission maxima at longer wavelengths (H1 and H3) continuously increased throughout the stationary phase. In contrast, the other humic-like fluorophore (H2) was relatively stable during the stationary phase (Student’s t-test, p > 0.05).

Fluorescence intensities of all fluorophores were comparable in the control and experimental treatments at the initiation of these treatments (Figures 3, 5). In the control treatment, the fluorescence intensities of all humic-like fluorophores remained unchanged during the incubation (Figure 5). Although the fluorescence intensities of protein-like fluorophores in the control treatment increased several fold during the 168-h incubation, these intensities were 1 order of magnitude lower than those observed for the experimental treatment at the end of the incubation (Figure 3).

A. macleodii has been reported to be a ubiquitous gammaproteobacterium in temperate oceans (López-Pérez et al., 2012). The possibility that A. macleodii substantially contributes to the consumption and alteration of labile DOM in surface waters of marine environments has also been considered (Tada et al., 2011; Pedler et al., 2014; Pedler Sherwood et al., 2015). Glucose was used as a labile substrate for A. macleodii incubation in this study. Previous studies demonstrated that coastal microbial communities completely consumed glucose within 2 days of incubation (Ogawa et al., 2001; Kawasaki and Benner, 2006). The decrease in the DOC concentration was coupled with the increase in A. macleodii abundance, indicating that the decrease in the DOC concentration during the exponential growth phase should be due to the consumption of glucose by the bacterial strain. Assuming that the glucose was completely consumed within several days of incubation, residual DOC was produced by A. macleodii.

Protein molecules that contain both tryptophan and tyrosine molecules usually show only tryptophan fluorescence because of energy transfer (Lakowicz, 2006). A previous study analyzed the fluorescence properties of surface DOM in the Sagami Bay and showed that only tryptophan-like fluorescence was evident in high-molecular-weight fractions, whereas tyrosine-like fluorescence was dominant in low-molecular-weight fractions (Yamashita and Tanoue, 2004). Thus, changes in the composition of protein-like fluorophores with A. macleodii growth possibly reflected the changes in the relative molecular weight of peptides/proteins, which were released and reused by A. macleodii. Tryptophan-like fluorophore clearly appeared at 24 h in our experiment (Figure 4), implying that large polymers, such as protein molecules, were predominantly released during the exponential growth phase. The incubations contained not only tryptophan-like fluorophore but also tyrosine-like fluorophore, which both increased during 24–72 h (Figure 4). This result suggested that large polymers were still released but that part of these exopolymers were degraded to small peptides. The decrease in tryptophan-like fluorophore was predominant, but tyrosine-like fluorophore was relatively stable at the latter part of the stationary phase (72–168 h), suggesting that the production of small peptides due to the degradation of large exopolymers was correlated with the reutilization of tyrosine-like fluorophores.

The reutilization of exopolymer by A. macleodii was suggested in the stationary phase while two humic-like fluorophores continued to accumulate, implying that unavailable DOM were produced from available substrate. The DOC concentrations were relatively stable during the stationary phase, as mentioned above. These results suggested that major portions of DOM produced by A. macleodii are unavailable to these bacteria, although a minor fraction of this DOM is reusable by the strain. The remaining DOC concentration produced by A. macleodii was estimated to be 41 ± 4 μM C by subtracting the DOC concentration at the end of control treatment (9.6 ± 1.5 μM C; Figure 1) from that of experimental treatment (51 ± 4 μM C; Figure 1). This remnant DOC concentration corresponded to approximately 4% of the initial glucose used for the A. macleodii incubation. Notably, this estimation is comparable to the results obtained by previous studies that conducted incubations of bacterial isolates or microbial communities with glucose. Gruber et al. (2006) showed that 3% of initial glucose was converted to residual DOC by the bacterial isolate P. chlororaphis during 36 days of incubation. In addition, 3–6% of initial glucose was found to convert to molecularly uncharacterizable DOM in marine microbial communities, and occurred after 90-day to 2-year periods of incubation (Ogawa et al., 2001; Shimotori et al., 2009; Koch et al., 2014). Such consistency in the production efficiency of residual DOC (i.e., 3–6% of initial substrate) suggests that neither microbial community structure nor microbial growth affected the efficiency of MCP.

The production of humic-like fluorophores has been observed in incubation experiments using marine microbial communities (Kramer and Herndl, 2004; Lønborg et al., 2009; Shimotori et al., 2009). This study demonstrated that humic-like fluorophores can be generated not only by microbial communities but also by single bacterial strains using simple substrates, such as glucose. This finding is consistent with those of Shimotori et al. (2012), who found that humic-like fluorophores were generated during incubations of several bacterial isolates in Marine Broth culture medium.

A humic-like fluorophore (H2) produced by A. macleodii was similar to a previously defined marine microbial humic-like peak M (Coble, 1996, 2007). This fluorophore also corresponded to a fluorophore that was found to be microbial recalcitrant FDOM with the time scale of the thermohaline circulation (Yamashita and Tanoue, 2008). Fluorescent components similar to H2 were obtained through parallel factor analysis (PARAFAC) of FDOM from the open ocean and were found to be related to apparent oxygen utilization in the deep ocean (Jørgensen et al., 2011; Tanaka et al., 2014), implying that H2 produced by A. macleodii is possibly recalcitrant DOM. On the other hand, peak M was produced by phytoplankton isolates (Romera-Castillo et al., 2010) and consumed partially by bacterial communities (Romera-Castillo et al., 2011). These results suggest that the reactivity of the peak M-type fluorophore was dependent on its source; specifically, the fluorophore produced by heterotrophic bacteria is possibly recalcitrant, while that produced by phytoplankton includes a labile fraction.

The other two humic-like fluorophores (H1 and H3), which peaked at longer emission wavelengths than that of H2, continued to be produced by A. macleodii during both the stationary phase and the exponential growth phase. Shimotori et al. (2009) also observed a fluorophore peaked at >500 nm emission during the incubation of coastal microbial communities with glucose. On the other hand, there was a high presence of PARAFAC components that were similar to a combination of H1 and H3 in humic acid fractions extracted from soils and sediments (He et al., 2006; Santín et al., 2009). These PARAFAC components were also considered to be terrigenous FDOM (Stedmon and Markager, 2005; Singh et al., 2013). These results suggest that the fluorophore that peaked at a longer emission wavelength (>500 nm) might be ubiquitously produced by heterotrophic bacteria but is enriched in humic acid fractions from soils and sediments. Notably, in Shimotori’s incubation studies (Shimotori et al., 2009), this fluorophore appeared in EEMs during the period of increasing bacterial abundance but disappeared from EEMs during the period of decreasing bacterial abundance. In the open ocean, this fluorophore is generally not present (Jørgensen et al., 2011; Tanaka et al., 2014). Thus, A. macleodii may be one of the key species that produces this fluorophore, and this fluorophore might be consumed by other microbes in natural environments.

The production rates of microbial humic-like fluorophore differ among substrates used in the incubation of marine bacterial communities (Jørgensen et al., 2014; Aparicio et al., 2015). For example, Aparicio et al. (2015) found that the production rates of marine humic-like fluorophore during the incubation of marine bacterial communities with Suwannee River humic acids were higher than those with glucose and acetate. Using different substrates for microbial community incubation reportedly cause changes in microbial community structure and/or bacterial physiology (Gómez-Consarnau et al., 2012). Therefore, in studies of bacterial communities, it is difficult to evaluate whether the factor regulating the production of humic-like fluorophores is bacterial species or physiology; in contrast, incubations using a bacterial isolate allow insight into the relationship between bacterial physiology and RDOM production.

The bacterial physiology was evaluated in terms of growth phases in this study. The quantity or quality of substrate is one of the critical factors in controlling bacterial growth (Jannasch and Egli, 1993). Therefore, the physiology of A. macleodii was likely altered by the change in substrate availability. The exponential growth phase of A. macleodii can be characterized as the period that the strain consumed glucose and produced various extracellular compounds, e.g., three humic-like fluorophores as well as a tryptophan-like fluorophore. The physiology of A. macleodii shifted to the stationary phase after the complete consumption of glucose, and the tryptophan-like fluorophore was consumed, but two humic-like fluorophores were produced. The results showed that the production mechanisms of humic-like fluorophores, which cannot be reused by the strain, are correlated with the physiology of A. macleodii. The humic-like fluorophore (H2) that is similar to recalcitrant microbial humic-like FDOM (Yamashita and Tanoue, 2008; Catalá et al., 2015) is only produced in the exponential phase, but other fluorophores (H1 and H3), which might be consumed by other microbes (Shimotori et al., 2009), are produced during both the exponential and the stationary phases of A. macleodii.

The relationship between bacterial growth and RDOM production that is apparent with A. macleodii was not consistent with the previous studies conducted by Gruber et al. (2006) and Eichinger et al. (2009). A. macleodii was found to produce extracellular humic-like fluorophores, which were unavailable to the strain during the exponential growth and the stationary phases. However, DOC produced by a different Alteromonas strain (A. infernus) was accumulated under conditions of carbon starvation of the isolate (i.e., the stationary phase in this study) during the incubation in which pyruvate was added every 48-hours as labile substrate (Eichinger et al., 2009). Specific compounds detected by mass spectrometry with electrospray ionization were produced in the exponential growth phase and remained at the end of P. chlororaphis incubation with glucose for 36 days, suggesting that recalcitrant compounds, which cannot be reused by the strain, were possibly produced during the exponential growth phase (Gruber et al., 2006). Although incubation conditions, such as temperature, the quality and quantity of substrate, and the composition of artificial seawater, varied among the studies, the results obtained by this study and the previous studies suggest that the physiology related to RDOM production might differ between bacterial species.