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Applied and Environmental Microbiology, June 2007, p. 3936-3944, Vol. 73, No. 12
0099-2240/07/$08.00+0     doi:10.1128/AEM.00592-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Aerobic Anoxygenic Phototrophic Bacteria Attached to Particles in Turbid Waters of the Delaware and Chesapeake Estuaries ^,

Lisa A. Waidner and David L. Kirchman^*

College of Marine and Earth Studies, University of Delaware, Lewes, Delaware 19958

Received 14 March 2007/ Accepted 23 April 2007

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Aerobic anoxygenic phototrophic (AAP) bacteria are photoheterotrophs that, if abundant, may be biogeochemically important in the oceans. We used epifluorescence microscopy and quantitative PCR (qPCR) to examine the abundance of these bacteria by enumerating cells with bacteriochlorophyll a (bChl a) and the light-reaction center gene pufM, respectively. In the surface waters of the Delaware estuary, AAP bacteria were abundant, comprising up to 34% of prokaryotes, although the percentage varied greatly with location and season. On average, AAP bacteria made up 12% of the community as measured by microscopy and 17% by qPCR. In the surface waters of the Chesapeake, AAP bacteria were less abundant, averaging 6% of prokaryotes. AAP bacterial abundance was significantly correlated with light attenuation (r = 0.50) and ammonium (r = 0.42) and nitrate (r = 0.71) concentrations. Often, bChl a-containing bacteria were mostly attached to particles (31 to 94% of total AAP bacteria), while usually 20% or less of total prokaryotes were associated with particles. Of the cells containing pufM, up to 87% were associated with particles, but the overall average of particle-attached cells was 15%. These data suggest that AAP bacteria are particularly competitive in these two estuaries, in part due to attachment to particles.

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Recent studies indicate that photoheterotrophic bacteria, which are capable of both phototrophy and heterotrophy, may be abundant and important in biogeochemical cycles of the oceans (3, 23, 28). One type of photoheterotroph, aerobic anoxygenic phototrophic (AAP) bacteria, can harvest light by use of bacteriochlorophyll a (bChl a) for the production of ATP (45). Phototrophy explains the higher growth rates (46) and viability (40) of these bacteria grown in the light compared with the results seen with dark-grown cultures. Some AAP bacteria reduce the production of photosynthetic pigments in response to higher organic carbon concentrations (27, 40). These data suggest that AAP bacteria would have competitive advantages over typical heterotrophs when organic carbon concentrations are low (23).

This hypothesis was supported by the discovery of AAP bacteria in the open ocean (28), where AAP bacteria can make up from <1% to 10% of the prokaryotic community (13, 18, 37). However, emerging evidence indicates that these bacteria may be as abundant in eutrophic as in oligotrophic environments (13, 37, 39). A global survey of bacteria containing bChl a and the reaction center pufM gene found that AAP bacteria were more abundant in the Long Island Sound and Chesapeake Bay than in the open ocean (37). Additionally, these bacteria were more abundant in the North Atlantic Ocean, where chlorophyll concentrations were higher, than in the North Pacific, where AAP bacteria comprised less than 5% of prokaryotes (13). These wide ranges suggest that more data on the abundance of AAP bacteria are needed to determine the ecological controls of these bacteria.

To explore what environmental factors control AAP bacteria, we enumerated cells containing bChl a by use of epifluorescence microscopy and pufM with quantitative PCR (qPCR) in samples from the Delaware and Chesapeake estuaries. Both estuaries are characterized by high concentrations of organic matter and nutrients and large inputs of terrestrial organic matter (21, 30). The Delaware estuary is mostly well mixed, with a large tidal influence from the adjacent North Atlantic Ocean (38). In mid-estuary, primary production is light limited due to high concentrations of suspended sediment (32). In contrast, the Chesapeake Bay is less influenced by tidal exchange with the North Atlantic Ocean (35), and suspended sediment concentrations at the turbidity maximum are about half those seen in the Delaware (36). We sampled well-oxygenated surface waters of the Delaware and Chesapeake estuaries where aerobic bacteria predominate, and we examined the abundance of AAP bacteria in particle-associated and free-living bacterial communities. In this study, AAP bacteria were abundant in turbid waters and were mostly associated with particles.

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Sampling and environmental parameters.
Samples were obtained on eight cruises during 2002 to 2006 from the main stems of the Delaware and Chesapeake estuaries at approximately 1-m depth. Light attenuation was measured using a Biospherical Instruments (San Diego, CA) light meter. In March 2005, turbidity was estimated by measuring optical backscatter with an OBS-3 instrument (D&A Instruments, Port Townsend, WA), and suspended solid (seston) concentrations were estimated from the optical backscatter data as described by Downing et al. (17). In March 2006, seston concentration (SSC) was calculated for the Delaware estuary using the following equation: SSC = 14.3k – 6.86, where k is the light attenuation coefficient (J. Sharp, unpublished data).

Bacterial production and chlorophyll a and nutrient concentrations were determined in whole-water samples only, as described previously (13). To estimate particle-attached AAP bacterial abundances, free-living microbes were removed from whole water by use of a modification of the method described by Crump et al. (16). In brief, the method consists of gentle reverse-gravity filtration through Whatman GF/D filters (nominal pore size, 2.7 µm). Magnetic-filter funnels (Pall Life Sciences) (47 mm) loaded with GF/D filters were floated in large beakers containing sample water. Water containing free-living bacteria flowed upwards through the filter, and this fraction was retrieved from the upper reservoirs by pipetting.

Microscopic detection of bChl a-containing cells.
Samples were preserved with 2% (final concentration) glutaraldehyde at 0 to 4°C for 1 to 4 h and then vacuum filtered (<20 lb/in²) onto 25-mm-diameter black 0.2-µm-pore-size polycarbonate filters. The filters were rinsed twice with 0.2-µm-filtered Milli-Q water, placed in cryovials, and held in liquid nitrogen or on dry ice (up to 8 days) before storage at –80°C. A section of the filter (approximate area, 61 mm²) was cut and stained on Parafilm in a 0.4 µg/ml solution of 4',6'-diamidino-2-phenylindole (DAPI) in 1x phosphate-buffered saline for 5 min. It was then dried on a 0.45-µm-pore-size filter (Millipore HA) mounted on a vacuum manifold, rinsed twice with filtered (pore size, 0.2 µm) Milli-Q water, and mounted on a slide with oil.

The procedure for counting total prokaryotes and AAP cells was previously described (13). Briefly, each of 30 fields of view was subjected to the following four exposures (excitation, emission): DAPI (360, 460 nm); bChl a (390, 750 nm); Chl a (480, 660 nm); and phycoerythrin (545, 610 nm). AAP bacteria were scored as DAPI and bChl a positive but chlorophyll a and phycoerythrin negative. Cells greater than 1.2 µm² in size were rejected from the bChl a-positive and CY3- and Chl a-negative masks. DAPI-positive cells were used to calculate total prokaryote counts per milliliter of sample water, and AAP cell numbers were calculated as a percentage of total DAPI-positive cells.

Quantitative PCR of pufM.
DNA from the August 2002, November 2002, and July 2004 samples was isolated from water prefiltered through polycarbonate filters (0.8 µm pore size) to minimize nonbacterial DNA numbers. All other samples either were subjected to gentle (reverse gravity) prefiltration through GF/D filters or were from whole water. Sample water (200 to 2,000 ml) was filtered onto 0.45-µm Durapore membranes (Millipore, Billerica, MA). The DNA on the filters was preserved in sucrose lysis buffer, extracted with phenol-chloroform, and further purified using an IsoQuick nucleic acid extraction kit (ISC Bioexpress, Kaysville, UT) or by cetyltrimethylammonium bromide extraction (14).

To target all riverine and estuarine pufM genes, we used a new primer pair for qPCR, pufM557F and pufM_WAWR. The pufM_WAW reverse primer (47) sequence exactly matches both Delaware River fosmid sequences 06H03 and 13D03 (43). We examined the target region of forward primer pufM557F (1) in 249 sequences, which included all available sequences from other environments (May 2006) and from 171 pufLM fragments generated by PCR amplification from bacterioplankton in the Delaware River (unpublished data). Of these sequences, less than 10% contained more than three mismatches or mismatches to the last five bases at the 3' end of the primer (see Table S1 in the supplemental material). This primer contained one mismatch at the 5' end to Del06H03 and two mismatches near the 3' end to Del13D03 (see Fig. S1 in the supplemental material).

Using Del06H03 as the standard for qPCR amplification, the target was detected at concentrations as low as two copies per reaction (data not shown). Amplification efficiencies of pufM qPCRs ranged from 82% to 88%, as determined by the slope of the regression of log copies with threshold cycle values. Detection of Del13D03, however, gave lower numbers. The amplification efficiency for this pufM sequence was 88%, but the target was detected at 200-fold-fewer copies than expected (data not shown). Therefore, Del06H03 was used as the standard for all qPCR assays.

The standard for qPCR was composed of a mixture of genomic and fosmid DNA from an uninduced culture of Escherichia coli EPI300 (Epicentre, Madison, WI) containing the Del06H03 clone. The final composition of the DNA included one copy of the fosmid molecule for every seven copies of the 16S rRNA gene of the host bacterium. Standards for qPCR included 10 to 10⁶ copies of pufM or 10³ to 10⁸ copies of the 16S rRNA gene.

The pufM gene was amplified under the following conditions: 10 min of denaturation and activation of the enzyme at 95°C followed by 40 cycles of denaturation at 95°C (15 s), annealing at 56°C (45 s), and extension and detection at 72°C (45 s). rRNA genes were amplified using the BACT1 primer pair described by Suzuki et al. (41) except that PCR products were detected with SYBR green I fluorescence. Cycling conditions for 16S rRNA genes were the same as for pufM qPCR except that the annealing temperature was 60°C and amplification was for only 30 cycles. Final concentrations of reagents in the amplification reaction mixtures were as follows: 1x Brilliant SYBR green Master Mix (Stratagene, La Jolla, CA), 80 pg/µl DNA, 0.096 µM each primer, and water to achieve a 12.5-µl final reaction volume.

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Direct enumeration of bacteriochlorophyll a-containing bacteria.
We examined the abundance of AAP bacteria by use of direct microscopic enumeration of bChl a-positive cells. AAP bacteria were very abundant in some samples from the Delaware estuary, reaching 34% of total prokaryotes (Table 1). In six transects of the Delaware estuary examined over 5 years, average prokaryote and AAP bacterial abundances ranged from 1.2 x 10⁶ to 3.2 x 10⁶ cells ml^–1 and 1.0x 10⁵ to 5.1 x 10⁵ cells ml^–1, respectively (Table 1). In these six transects, the overall mean (± standard error [SE]) AAP bacterial abundance was 12% ± 8% of prokaryotes. Total prokaryote abundances decreased as salinity increased through the estuary, but there was no consistent trend in the spatial variation in AAP bacterial abundances (data not shown). While AAP cells varied in abundance from 1% to 34% of total prokaryotes, water column turbidity also varied greatly. The light attenuation coefficient ranged from 0.6 to 5.8 m^–1, and the overall average was 2.3 ± 1.3 m^–1 (Table 1).

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TABLE 1. Average (±SE) prokaryote and AAP bacterial abundances in the Delaware estuarya

In all five transects of the Delaware estuary and two of the Chesapeake, there was a positive correlation between relative AAP bacterial abundance (in percentages of total prokaryotes) and light attenuation (r = 0.53 [P < 0.001]; n = 69) (Fig. 1). In the Delaware alone, relative AAP bacterial abundance was positively correlated with light attenuation (r = 0.48 [P < 0.001]; n = 51), but in the Chesapeake there was no significant relationship (r = 0.37 [P > 0.05]; n = 18). Light attenuation coefficients in the Chesapeake were lower than in the Delaware, ranging from 0.5 to 1.6 m^–1 (Fig. 1). Additionally, the average AAP bacterial abundance in the Chesapeake was 6% ± 5% of prokaryotes, twofold lower than observed in the Delaware. In contrast to relative AAP bacterial abundance results, total prokaryote abundance did not covary significantly with light attenuation in either estuary (Delaware, r = 0.08 [P > 0.05]; n = 51; Chesapeake, r = 0.20 [P > 0.05]; n = 18).

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FIG. 1. Relationship of relative bChl a-containing cell abundance (percentage of total prokaryotes) to light attenuation in the Delaware and Chesapeake estuaries. The error bars represent averages for all data points. Error values represent standard errors of the linear regression of light attenuation with depth and for 30 fields of view (bChl a).

In addition to light attenuation, we examined the effects of other parameters on the distribution of AAP bacteria in the Delaware estuary (Table 2). Relative AAP bacterial abundance significantly covaried with concentrations of ammonium and nitrate plus nitrite (Table 2). There was also a positive correlation between relative AAP bacterial abundance (percentage of total prokaryotes) and total prokaryote abundance. Notably, however, total prokaryote abundance was not significantly correlated with any parameter except bacterial production as measured by thymidine incorporation (Table 2). In the Chesapeake Bay, abundance of AAP bacteria did not significantly covary with nitrate (r = 0.6 [P > 0.05]; n = 8) or ammonium (r = 0.01 [P > 0.05]; n = 8) concentrations or with total prokaryote abundance (r = 0.07 [P > 0.05]; n = 8).

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TABLE 2. Correlation of total and AAP bacterial abundances in whole water in the Delaware estuarya

Particle-attached and free-living AAP bacteria.
To test methods for separating free-living from particle-attached bacteria, we compared AAP bacterial abundances in whole water and the isolated free-living fraction by vacuum and reverse-gravity filtration. In 8 of 10 samples, the free-living AAP bacterial abundance was lower with the gentle GF/D treatment than with the vacuum polycarbonate treatment regardless of polycarbonate filter pore size (Table 3). In two of four samples subjected to reverse-gravity filtration, free-living AAP bacterial abundance in the polycarbonate filtrate was higher than that seen in the GF/D filtrate results (Table 3). These data indicate that, in most cases, gentle prefiltration through GF/D filters was best suited for separating free-living and particle-attached bacteria.

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TABLE 3. Comparison of two methods for separating free-living and particle-attached prokaryotes and AAP bacteria

We estimated the abundance of particle-attached bacteria by subtracting values for the free-living fraction from those for the corresponding whole water. In two transects of the Delaware estuary, most AAP bacteria were attached to particles, while most total prokaryotes were free living (Table 4). In March 2005, the particle-attached fraction contained 41% to 94% of all AAP bacteria (Table 4). In contrast, most total prokaryotes were in the free-living fraction, with only 2.5% to 28% of prokaryotes in the particle-attached fraction, except at the mouth of the bay, where 47% of prokaryotes were associated with particles. Total AAP bacterial abundance was lower in March 2006 than in March 2005 (Table 1), but the particle-attached community accounted for 31% to 92% of all AAP bacteria (Table 4). Again, as in March 2005, while AAP bacteria were mostly in the particle-attached fraction, most of total prokaryotes were free living (90% or higher for all samples).

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TABLE 4. Average percentages of cells associated with particles in the Delaware estuary

For five of nine samples from March 2006, the values for the estimates of abundance of total prokaryotes attached to particles were negative (Table 4), because the estimated abundance in whole water was lower than the abundance for the free-living fraction. Of these five samples, three gave values statistically significantly less than 0 (t test; P < 0.05). The negative values were most likely due to particulate material obscuring cells, leading to underestimations of total prokaryote numbers.

To further examine abundance in both fractions, we used a qPCR assay for the pufM gene, a marker for AAP bacteria (18, 37). Both qPCR and microscopy estimates indicated high AAP bacterial abundances in the particle-attached community in March 2005 (Fig. 2). For all samples, microscopy abundance estimates of AAP bacteria in whole water exceeded those determined for the free-living fraction (Fig. 2A). For three samples, qPCR estimates yielded higher abundances of pufM-containing bacteria in the free-living fraction than in the whole water, resulting in negative values for particle-attached AAP bacterial abundances (Fig. 2B). For the remaining five samples, however, particle-associated AAP bacterial abundances ranged from 26 to 87% of total AAP bacteria. Overall, the pufM data suggest that 2 to 50% of bacteria were AAP bacteria (Fig. 2B), assuming two 16S rRNA gene copies per bacterial cell (19).

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FIG. 2. Relative abundance of bChl a- and pufM-containing bacteria in whole water and in the free-living fraction in the Delaware estuary in March 2005. Abundances were measured in whole-water samples (solid black bars) and in the free-living fraction (size, <3 µm) (white bars). (A) Values for abundance of AAP bacteria are expressed as percentages of total prokaryotes. Error bars represent the standard errors for 30 fields of view. Measurement of water column turbidity was plotted as optical backscatter (solid line). (B) Copy numbers of the pufM gene were normalized to numbers determined for the 16S rRNA gene. Error bars represent the standard errors for the results of four qPCRs. The figure legend refers to both panels.

The abundance of particle-associated AAP bacteria was again high in March 2006, as measured by both direct counts and qPCR (Fig. 3). In seven of nine samples, whole-water pufM bacterial abundances exceeded those determined for the free-living fraction, and 17% to 83% of bacteria with pufM were estimated to be associated with particles. Particle-attached AAP bacteria as determined by qPCR were most abundant just downstream of the turbidity maximum, approximately 60 to 80 km from the mouth (Fig. 3; Table 4). Abundances of free-living cells containing pufM followed the trends of free-living bChl a-containing cells, and estimates by both qPCR and microscopy indicated that AAP bacteria comprised 0.4% to 4.5% of bacteria in the free-living fraction (Fig. 3A and B). For the whole water, estimates of numbers of pufM-containing bacteria ranged from 0.6% to 4.4% of bacteria, while microscopy estimates of AAP bacteria gave values as high as 14% of prokaryotes.

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FIG. 3. Relative abundance of bChl a- and pufM-containing bacteria in whole water and in the free-living fraction in the Delaware estuary in March 2006. Abundances were measured in whole-water samples (solid black bars) and in the free-living fraction (size, <3 µm) (white bars). (A) Values for abundances of AAP bacteria are expressed as percentages of total prokaryotes. Error bars represent the standard errors for 30 fields of view. (B) Copy numbers of the pufM gene were normalized to numbers determined for the 16S rRNA gene. Error bars represent the standard errors for the results of four qPCRs. Water column turbidity was plotted as an attenuation coefficient (solid line).

To examine the effect of turbidity on particle attachment of AAP bacteria, we examined the relationship between particle-attached AAP abundance and calculated seston concentration (Fig. 4). Combining all the data from both methods for determining AAP bacterial abundance, the relationship between particle-attached abundance and seston concentrations was significant (r = 0.44 [P < 0.05]; n = 31). Both microscopy (r = 0.65 [P < 0.01]; n = 16) and qPCR (r = 0.68 [P < 0.01]; n = 15) abundance estimates for particle-associated AAP bacteria were positively correlated with seston concentrations (Fig. 4).

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FIG. 4. Particle-associated bChl a- and pufM-containing cell numbers as a function of suspended sediment load. Seston concentrations were calculated in March 2005 from optical backscatter and in March 2006 from light attenuation. Particle-associated pufM- and bChl a-containing cell numbers were obtained by subtracting free-living from whole-water values.

Abundance estimates by qPCR and microscopy.
To examine in more detail the abundance estimates obtained with qPCR and microscopy, we enumerated bChl a- and pufM-containing bacteria in additional transects of the Delaware estuary in 2002 and 2004 (Fig. 5). In these samples, the microscopy samples were from whole water, but the DNA used for pufM analyses was isolated from water prefiltered with a <0.8-µm-pore-size filter to minimize eukaryotic DNA. The microscopy abundance estimates were higher than the estimates determined by pufM qPCR. Assuming two copies of 16S rRNA genes per bacterium (8, 19), the AAP abundances calculated for these three transects were from <1% to 4% of bacteria. In contrast, the bChl a-containing cell numbers for these samples ranged from 1% to >30% of prokaryotes (Fig. 5).

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FIG. 5. bChl a-positive cells in whole-water samples and pufM quantities in DNA from the <0.8-µm-size fraction. Error bars represent the standard errors for 30 fields of view. Copy numbers of pufM were normalized to copy numbers for the 16S rRNA gene. Error bars represent the standard errors for four qPCRs. General trends determined by locally weighted scatterplot smoothing lines were plotted for bChl a cells (solid line) and pufM per 16S rRNA gene (dashed line). Transects of the Delaware estuary represent values obtained for August 2002 (A), November 2002 (B), and July 2004 (C).

There were also differences in spatial abundance patterns as estimated by the two methods, resulting in different correlations between estimates of AAP bacteria by microscopy and by qPCR (Fig. 5). In August 2002, there was no correlation between direct counts and pufM abundances (r = 0.13 [P > 0.05]; Fig. 5A). In November 2002, there was a negative relationship (r = 0.67 [P < 0.05]; Fig. 5B). In contrast, in July 2004, there was a positive correlation between pufM abundances and bChl a-containing cell results (r = 0.82 [P < 0.05]; Fig. 5C).

In March 2005 and March 2006, we directly compared abundance estimates calculated using both methods, assuming two copies of the 16S rRNA genes per bacterial cell and one pufM per AAP bacterial cell (Fig. 6). Out of 17 samples, 8 whole-water samples were close to the 1:1 line (Fig. 6A), corresponding to a ratio of one pufM-containing cell to one bChl a-containing cell (pufM-to-bChl a ratio). The mean (±SE) ratio in March 2006 was 0.32 ± 0.05, indicating an underestimation of AAP bacteria determined by qPCR compared to microscopy estimates. In contrast, in March 2005, most ratios were between the 1:1 and 4:1 lines, and the average pufM-to-bChl a ratio was 2.2 ± 0.4 for whole-water results. Overall, the average (±SE) ratio for both years was 1.2 ± 0.4 for whole water (n = 17), indicating that the two methods yielded similar estimates of AAP bacterial abundances in whole-water samples.

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FIG. 6. Percentages of bacteria containing bChl a and pufM in whole water (A) and the free-living fraction (B). The percentages of prokaryotes with pufM were calculated as follows: (pufM copies/2 copies of 16S rRNA gene) x 100. Lines (1:1 and 4:1) are the ratios of pufM-containing cells to bChl a-containing cells.

In the free-living samples, few ratios were near the 1:1 line (Fig. 6B). The mean ratio (±SE) in March 2005 for free-living samples was 8.3 ± 2.0. In contrast, in March 2006, AAP bacteria were underestimated by qPCR compared to microscopy estimates (Fig. 6B), with a mean (±SE) pufM-to-bChl a ratio of 0.50 ± 0.12. Overall, in both years, the mean ratio (±SE) was 4.2 ± 1.6 for the free-living fraction. These data suggest that not all pufM-containing cells were detected by microscopy, especially those AAP cells in the free-living fraction.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The aim of this study was to examine the relative abundance and distribution of AAP bacteria in two estuaries in order to elucidate the factors controlling these potentially important photoheterotrophic bacteria. It was first thought that AAP bacteria would be more competitive in oligotrophic waters than in eutrophic environments (27), since these bacteria can supplement ATP production via photophosphorylation (45). This hypothesis was supported by studies of freshwater and marine AAP bacteria (5, 40) as well as by reports of high abundances in oligotrophic oceans (27-29). However, more recent data suggest the opposite of the original hypothesis. AAP bacterial abundance has been found to be higher in estuarine, coastal, and shelf waters than in some oligotrophic oceans (37, 39, 48). One explanation for these conflicting results could be the high diversity of AAP bacteria. Ecotypes of AAP bacteria in estuaries may respond to environmental factors differently than their open-ocean counterparts. Our data indicated that inorganic nitrogen concentrations positively influenced the relative abundance of AAP bacteria but did not contribute to the success of the prokaryote community as a whole. However, nutrient levels only explained part of the variation in AAP bacterial abundances. Among several other factors, light may be important in determining the success of AAP bacteria (40, 45).

Since light provides an energy advantage to these bacteria, it is not surprising that AAP bacteria are most abundant in the euphotic zones of the oceans (13, 27, 39). In this study, however, AAP bacterial abundance was inversely correlated with light availability. One explanation for these data is that low light levels lead to high cellular concentrations of bChl a (12, 25, 46) and thus higher detection by infrared epifluorescence microscopy. However, the qPCR data also indicated higher abundance of AAP bacteria in waters with higher light attenuation. Light attenuation is due to a combination of factors, including the presence of colored dissolved organic matter (9, 11) and suspended particles (6, 7). In the Baltic, bChl a and colored dissolved organic matter concentrations were positively correlated (26), suggesting an inverse relationship of AAP bacterial abundance with light. In the Delaware, however, the negative correlation with light is consistent with our observation of an association of AAP bacteria with particles. This relationship with particles may also explain the abundance of AAP bacteria in the Yangtze River estuary (18), where concentrations of suspended particles are very high (44). The preference of AAP bacteria for particles may also be important in open-ocean regimens. In metagenomic clones from the Sargasso Sea, the estimated AAP abundance in free-living bacterioplankton (<0.8 µm) was twofold lower than that of the 3- to 20-µm fraction (48), suggesting that AAP bacteria are associated with marine snow in oligotrophic waters.

We examined AAP bacteria in the free-living community by separating them from attached bacteria via reverse-gravity filtration through GF/D filters. Several observations indicated that free-living AAP bacteria were not trapped by the GF/D filters (nominal pore size, 2.7 µm) in the gentle prefiltration process. First, an examination of microscope images confirmed that many of these bacteria were attached to particles, whereas most other prokaryotes were in the free-living fraction (see Fig. S2 in the supplemental material). Second, even though all cells were physically treated the same, most total prokaryotes were free living, but 50% or less of AAP cells were in this fraction. Finally, an examination of cell sizes calculated from 1,260 fields of view indicates that the maximum lengths and widths of all AAP cells were less than the pore size of the GF/D filters (see Table S2 in the supplemental material).

Evidence from studies of cultured AAP bacteria is consistent with our finding of particle attachment by this group of bacteria. Dinoroseobacter shibae, an AAP bacterium isolated from dinoflagellates, produces quorum-sensing autoinducers (42), which may assist these bacteria in up-regulating hydrolytic enzymes or antibiotics in order to more efficiently colonize or use organic matter (10, 20). Porphyrobacter tepidarius, a member of the Sphingomonadales subgroup of Alphaproteobacteria, and two uncharacterized strains of AAP bacteria form extracellular matrices (45), which may facilitate attachment to particles.

The reduced oxygen concentration in particles may be important to the success of particle-associated AAP bacteria. In this microenvironment, low oxygen may reduce respiration by all aerobes, while photophosphorylation by AAP bacteria would be unaffected. In fact, Hoeflea phototrophica, a marine AAP bacterium isolated from dinoflagellates, is capable of growth with temporary reductions in oxygen concentration (4). It is not clear whether this growth is due to increased light-driven ATP production, but reduced oxygen does stimulate the synthesis of bChl a and reaction center proteins in freshwater and marine isolates (5, 40, 45). However, complete removal of oxygen halts growth and photosynthetic electron transfer in cultivated AAP bacteria (45).

Several lines of evidence indicate the bacteria we detected on particles were not obligate anaerobes. The surface waters of the Delaware and Chesapeake estuaries are well oxygenated (31, 38). Although the particles and aggregates in these waters may have low oxygen concentrations, they are most likely not anoxic. Only the largest particles of marine snow (>500 µm) are anoxic for a short time (33), and most particles are only partially depleted of oxygen in the dark (2). In this study, the particles most likely containing attached anaerobes in the estuary would be in the riverine portion of the Delaware estuary. However, genomic DNA from two types of AAP bacteria from this region of the estuary contains genes indicative of aerobiosis (43).

While previous studies examined AAP bacterial abundances in whole water by microscopy (13, 37, 39), and in the free-living fraction by qPCR (18, 37), this study was the first to examine both fractions by both methods. The two methods differ in that one relies on expression and the other on the presence of genes encoded by the puf superoperon. In some samples, qPCR estimates were less than those determined by microscopy. One explanation could be artificially high estimates of 16S rRNA gene copies due to amplification of plastid genes. All of the underestimated values were from March 2006, when 16S rRNA gene copy numbers per total DNA were high (see Table S3 in the supplemental material). Although the 16S rRNA gene primer pair was designed specifically for bacteria, some amplification of plastid DNA is possible (M. T. Suzuki, personal communication). Another explanation is that not all phylotypes of pufM were efficiently amplified with the 557F-WAWR primer pair. The reverse primer was designed to amplify most known environmental pufM sequences (47), but the forward primer may miss at least 10% of pufM sequences (37, 47). For all whole-water samples, however, the estimates determined by qPCR were statistically equal to those determined by microscopy (Student's t test [P > 0.05]; n = 17). This was not the case for the free-living fraction results (Student's t test [P < 0.05]; n = 17).

Microscopy appeared not to detect all free-living AAP bacteria estimated to be present by qPCR. For 35% of the whole-water samples and 41% of the free-living fraction samples, qPCR abundance estimates were significantly greater than microscopy estimates (Student's t test [P < 0.05]). One explanation is that not all cells were detectable by microscopy because of low cellular bChl a concentrations, particularly in the free-living fraction. Among other factors, light and oxygen levels could lead to different bChl a cellular levels in free-living and particle-attached AAP bacteria. Indeed, decreased light and oxygen levels increase bChl a synthesis rates in AAP bacterial cultures (40, 46).

We determined that numbers of AAP bacteria were inversely correlated with light availability and that this relationship was partly explained by the association of these bacteria with particles. This observation has implications for the ecology of these photoheterotrophs and their impact on aquatic food webs. AAP and other bacteria associated with particles can have advantages over planktonic cells, since they may be less susceptible to grazing than those in the free-living fraction (22), are physically associated with rich sources of dissolved and particulate organic matter (15, 34), and are thus capable of faster growth than their free-living counterparts (16, 24). Because of their association with particles, attached AAP bacteria may be important for the dynamics of the bacterial community and higher trophic levels. Additionally, since AAP bacteria are capable of obtaining extra energy from light, their contribution to organic matter consumption may be even greater than their abundances would suggest. A better understanding of the photoheterotrophic lifestyle is important for our understanding of matter and energy fluxes in aquatic environments.

 
We thank Matthew Cottrell for his support and advice for enumeration of AAP bacteria. We also thank Vanessa Michelou, Hila Elifantz, Ogugua Anene-Maidoh, and the captain and crew of the R/V Cape Henlopen and R/V Hugh R. Sharp for assistance with sample collection. Chris Sommerfield, Jon Sharp, and Eric Wommack provided valuable support as chief scientists of cruises in the Delaware and Chesapeake estuaries. We thank Chris Sommerfield for optical backscatter data and Jon Sharp and Feng Chen for nutrient data. Marcelino Suzuki, Mike Schwalbach, and Natalya Yutin graciously provided insight into optimization of pufM quantitative PCR.

This work was supported by DOE grant DE-FG02-97ER62479 and NSF grant MCB-0453993.

 
* Corresponding author. Mailing address: College of Marine and Earth Studies, University of Delaware, Lewes, DE 19958. Phone: (302) 645-4375. Fax: (302) 645-4028. E-mail: kirchman{at}udel.edu

Published ahead of print on 27 April 2007.

Supplemental material for this article may be found at http://aem.asm.org/.

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Applied and Environmental Microbiology, June 2007, p. 3936-3944, Vol. 73, No. 12
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