Distribution and Activity of Bacteria in the Headwaters of the Rhode River Estuary, Maryland, USA
By: Parke A. Rublee, Susan M. Merkel, Maria A. Faust, and Joseph Miklas
Rublee, P.A. S.M. Merkel, M.A. Faust, and J. Miklas. 1984. Distribution and activity of bacteria in the
headwaters of the Rhode River estuary, Maryland, USA. Microbial Ecology 10:243-255.
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A transect along the axis of the headwaters of a tidal estuary was sampled for microbial, nutrient, and physical
parameters. Chlorophyll a averaged 42 μg 1-1 and phytoplankton comprised an estimated 80% of the total
microbial biomass as determined by adenosine triphosphate (ATP). Bacterial concentrations ranged from 0.3-
53.9 × 106 cells ml-1 and comprised about 4% of the total living microbial biomass. Bacterial production,
determined by 3H-methyl-thymidine incorporation was about 0.05-2.09 × 109 cells 1-1 h-1, with specific growth
rates of 0.26-1.69 d-1. Most bacterial production was retained on 0.2 μm pore size filters, but passed through 1.0
μm filters. Significant positive correlations were found between all biomass measures and most nutrient
measures with the exception of dissolved inorganic nitrogen nutrients where correlations were negative.
Seasonal variability was evident in all parameters and variability among the stations was evident in most. The
results suggest that bacterial production requires a significant carbon input, likely derived from autotrophic
production, and that microbial trophic interactions are important.
Coastal and estuarine systems are sites where competition for nutrients by microorganisms and remineralization
of photosynthetically fixed carbon are important processes. High rates of bacterial activity have been found in
such systems for both inorganic nutrients and organic compounds [10, 29]. For example, Faust and Correll [10,
11] measured the uptake of orthophosphate by 2 size fractions of suspended plankton in the Rhode River over a
period of 12 months. Except for summer months, the uptake of orthophosphate was predominantly by the
smallest size fraction ( < 5.0 μm), which they observed was primarily bacteria. Generally, only 1-6% of the
uptake was by the larger size fraction (> 5.0 μm), composed primarily of phytoplankton. Palumbo  found
rapid turnover times (1-2 days) but a high degree of variability in kinetic parameters of bacterial uptake of
mixed amino acids in the Newport River estuary, North Carolina. Salinity, temperature, heterotrophic biomass,
and supply of organic material all seemed to be important factors in regulating the bacterial activity.
Utilization of carbon fixed by primary production is a primary role of heterotrophic microorganisms, and recent
studies suggest that the magnitude of this role has been underestimated [26, 36]. Williams  used a model to
suggest that bacteria may utilize as much as 50% of the primary production of phytoplankton in coastal areas,
and that this magnitude of uptake is compatible with current trophic models. Similarly, Nixon  has pointed
out that earlier conceptualizations of remineralization pathways did not include bacteria as major contributors to
that process, but that recent experimental work has pointed to its fundamental importance in some systems. The
sources of dissolved organic carbon for bacterial uptake include excretion by primary producers, "sloppy
eating" by zooplankters, and zooplankton fecal material. The fate of this carbon material is not only
mineralization to inorganic carbon, but also production of bacterial biomass. Several authors [2, 36] have
suggested the trophic importance of bacteria as a link between phytoplankton and higher consumers.
Estuarine systems may exhibit even greater dependence upon trophic links through microheterotrophs than
oceanic systems since detrital pathways in estuarine systems are frequently well developed, especially where
surrounding marshes are extensive . In these systems, important interactive mechanisms for microbial
activity include not only the detrital pathway but also bacteria-particle relationships.
In this study we have utilized acridine oragne direct counts (AODC) and 3H-thymidine incorporation to
determine bacterial abundance and activity in the headwaters of a brackish tidal estuary, the Rhode River, in
Maryland. We have also assessed some of the physical and chemical factors that may be important in
determining bacterial distribution, activity, and the importance of these organisms to nutrient flux in the Rhode
Materials and Methods
Samples were collected at 7 stations in the Rhode River, a subestuary of Chesapeake Bay, on the east coast of
North America (38°51'N, 76°32'W). Five of these stations were along a transect which ran up the axis of the
river and headwater creeks (Fig. 1). The transect began in the river at station 4, continued upstream past a
mudflat, station 5, and into a tidal creek, Muddy Creek, surrounded by an irregularly flooded Typha angustifolia
marsh, station 6. The transect continued up 2 tributaries of Muddy Creek, Main Branch and North Branch.
Stations 7 and 8 are located in these 2 branches, respectively, at the position where there is a transition from
marsh to forested wetland. Salinity along the transect ranged from 18 ppt at station 4 during the summer to 0 ppt
at stations 7 and 8 during spring runoff periods. Although there is a tidal influence in the Rhode River
(approximately 35 cm), regional meteorological events such as wind and changes in barometric pressure can
cause changes in water level of over 1 m.
Two additional stations were sampled, a dock station and a low marsh tidal flume station. The dock station is
located near station 4 (Fig. 1) and was sampled on only 1 day, primarily to examine the size distribution of
heterotrophic activity. The low marsh station, located close to station 6, was sampled on 11 tidal cycles. Results
of the tidal cycle studies are given elsewhere , but we have utilized some of the data on bacterial
distribution and activity derived from that study to complement our observations from the other stations.
The transect was sampled 18 times at 2-6 week intervals at high tide. Water for nutrient analyses was collected
continuously as the boat moved from one station to the next. Water was pumped through a peristaltic pump into
5 gallon acid-washed polyethylene jugs, one for each segment of the transect. Microbial parameters and
dissolved organic compounds were determined from surface samples taken by immersing 1 gallon polyethylene
bottles just below the water surface. Subsamples for bacterial enumeration were fixed in the field with
formaldehyde to a final concentration of 2% (v/v). Salinity and temperature were measured at each station with
a Beckman RS5-3 salinometer, and dissolved oxygen was measured with a YSI Model 54 oxygen meter. All
samples were returned to the laboratory within several hours, and microbial samples were kept on ice.
Concentrations of organic matter, nitrogen, and phosphorus were determined from the integrated samples. Total
and dissolved organic matter were measured as chemical oxygen demand (COD) before and after filtration
through 0.4 μm membrane filters . HgSO4 was added to these samples prior to analysis to eliminate chloride
interference . Nitrate and ammonia were determined by a reduction and oxidation, respectively, to nitrite,
followed by nitrite analysis . Total phosphorus was determined by digestion with perchloric acid and
measured as orthophosphate by the stannous chloride method [1, 20]. Total particulate phosphorus was
calculated as the difference between total phosphorus and total dissolved phosphorus. Dissolved organic
phosphorus was calculated as the difference between dissolved orthophosphate and dissolved total phosphorus.
Total particulates were determined by weight after filtration onto Nuclepore filters (0.4 μm pore).
Various microbial parameters were determined from the surface samples. Bacterial concentrations were
determined using the acridine orange direct count method (AODC) of Hobbie et al. . Chlorophyll a was
determined by sample filtration through a glass fiber filter followed by maceration and an acetone/DMSO
extraction . Absorption of the extract was determined at 663 nm before and after acidification with 1.0 N
HCl, and pigment content determined by the equations given in Standard Methods . Heterotrophic activity
was measured as 3H-methyl thymidine incorporation following the method of Fuhrman and Azam [13, 14].
Briefly, 5 ml samples were incubated with 1 μCi of labeled thymidine (20 Ci/mmol-1, 1 nM final concentration)
for 30 min at room temperature and then filtered onto a 0.2 μm pore size polycarbonate membrane filter.
Incorporation of label was determined by liquid scintillation counting after extraction with trichloroacetic acid
as recommended by Fuhrman and Azam [13, 14] and correction for quenching by the channels ratio method.
All values were corrected to in situ temperature by assuming a Q10 factor of 2. Occasionally additional
replicates were incubated for determination of the size partitioning of thymidine incorporation. Following
incubation, replicates were filtered through 1.0, 3.0, or 5.0 μm pore Nuclepore membrane filters. Adenosine
triphosphate (ATP) concentration was determined following extraction in a boiling Tris buffer by the method of
Holm-Hansen and Booth 1181. Electron transport system (ETS) activity was measured by a tetrazolium
reduction method 1281. Dissolved primary amines , dissolved monosaccharides 1191, and dissolved total
carbohydrates  were also determined from surface samples.
Physical and chemical characteristics of the study site varied both seasonally and by station. Overall, salinity
ranged from 0-17.6 ppt (mean = 8.4 ppt, SE = 0.6, n = 86). Both station and sampling date contributed
significantly to the variability in salinity (ANOVA, F = 33.5, P < 0.01), and Duncan's multiple range test
separated the stations into 2 groups. Stations 6, 7, and 8 comprised one group with a lower average salinity, and
the downstream stations 4 and 5 comprised the second group. Salinity was occasionally high, however, even at
the upstream stations during 1980 and 1981 which were drought years. pH ranged from 6.1-9.0 (mean = 7.5, SE
= 0.06, n = 80) during the course of the study, and lowest values were found at the upstream stations. Mean total
suspended particle concentration was 24.7 mg 1-1 (SE = 2.0, n = 85), with a range from 4.7-108.5 mg 1-1.
Temperature was generally higher at the downstream stations. Analysis of variance indicated that significant
differences occurred among stations and sampling dates for total suspended particulates (F = 3.8, P < 0.01), pH
(F = 34.9, P < 0.01), and temperature (F = 54.7, P < 0.01). Mean values for these parameters by station are
given in Table 1.
Nutrient concentrations were also variable. All nutrient parameters had significant seasonal variation at the P <
0.05 level or better based on analyses of variance. Analyses of variance also indicated significant differences
among stations for dissolved nitrogen nutrients and orthophosphate. Mean values for nutrient concentrations at
stations are given in Table 2.
The concentration of bacteria ranged from 0.3-53.9 × 106 cells ml-1, with a mean value of 7.3 × 106 cells ml-1
(SE = 0.98 × 106 cells, n = 86). An analysis of variance indicated that there was no significant difference in
bacteria concentration among stations 4 through 8 of the transect (F = 1.11, P = 0.37). Seasonal patterns were
evident in bacterial concentrations, with fall peaks in bacteria in both 1980 and 1981 (Fig. 2).
Incorporation of 3H-thymidine followed a pattern similar to that of bacterial concentrations (Fig. 3). Mean
incorporation of thymidine over the 30 min incubation period was 2.89 x 104 dpm 5 ml-1 (SE = 0.45 × 104 dpm,
n = 60), with a range from 0.01-17.89 × 104 dpm 5 ml-1. There was no difference among stations (ANOVA, F =
2.00, P = 0.16) but a strong seasonality was evident (ANOVA, F = 8.92, P < 0.01). Thymidine incorporation
was found primarily in the smallest size fractions on 4 dates when replicate samples were filtered through
different pore size filters (Table 3). Generally, 60-70% of the incorporated thymidine was found in the 0.2 μm-
1.0 μm size fraction, and smaller percentages were found in larger size fractions up to >3.0 μm (Table 3).
Chlorophyll a distribution was similar to that of bacteria, except that maximum values were found in the fall of
1981 rather than 1980, although peaks in concentration were evident in both years (Fig. 4). Mean chlorophyll a
concentration for the data set was 42.1 μg 1-1 (SE = 5.9 μg, n = 84), with a range from 0.5-244.7 μg 1-1.
Variability in chlorophyll a concentration was attributable to both station and sampling date (ANOVA, F =
10.6, P < 0.01). Duncan's multiple range test found 2 groups, with stations 4, 5, 6, and 7 in the first group, and
stations 5, 6, 7, and 8 in the second group. In general, the highest concentrations of chlorophyll a were found at
ATP concentrations ranged from 0.3-43.1 μg ATP 1-1, with a mean value of 5.4 μg ATP 1-1 (SE = 1.2 μg, n =
50). The highest ATP values were generally found at the upstream stations. Similarly, values for ETS, the
community respiration measure, were highest at the upstream stations and had a highly significant positive
correlation with ATP (r = 0.94, P < 0.01, n = 39). Mean ETS value was 26.02 μg O 1-1 (SE = 7.1 μg, n = 42)
with a range from 0-251.2 μg O2 1-1 h-1. Strong positive correlations were found between chlorophyll a and both
ATP (r = 0.929, P < 0.01, n = 48) and ETS (r = 0.885, P < 0.01, n = 40); correlations with bacteria were not as
large for ATP (r = 0.453, P < 0.01, n = 50) or ETS (r = 0.308, P < 0.05, n = 42).
Significant correlation coefficients were found between chlorophyll a and many of the nutrient parameters
(Table 4). Dissolved nitrogen components exhibited negative correlations with chlorophyll a, while dissolved
phosphorus components exhibited positive relationships with chlorophyll a. Positive correlations were found
between chlorophyll a and most particulate nutrient parameters. Similar significant correlations were found
between bacteria and the nutrient parameters (Table 4).
Heterotrophic activity correlated significantly with chlorophyll a and with bacterial concentration in the overall
data set (Table 5). However, if the correlations were determined by station, a different pattern emerged.
Significant correlations between 3H-thymidine incorporation and bacterial concentration were found only in the
upstream stations 6, 7, and 8 (Table 5). Similarly, the only significant correlation between chlorophyll a and
thymidine incorporation occurred at station 8.
Microbial distribution and dynamics in the Rhode River estuary exhibit some similarities to other estuarine
systems. For example, bacterial concentrations are similar to those found for many estuarine and nearshore
systems [3, 4, 9, 21, 24, 25, 29]. The seasonal distribution of bacteria was also typical. Warmer summer
temperatures foster larger bacterial populations [e.g., 29]. The spatial distribution of bacteria was not typical,
however, as there was a significant positive correlation of bacterial numbers with salinity (r = 0.446, P < 0.01, n
= 86). In most studies of estuarine systems, the numbers decrease with increasing salinity [e.g., 3, 29]. Ducklow
 however, has documented a destratification event in the York River, Virginia that resulted in a doubling of
bacterial concentrations although salinity changed less than 2 ppt. It may be that the complexity of our estuarine
system, that is, high variability in salinity, nutrient concentrations, and freshwater inputs as well as the
variability resulting from meteorological conditions either obscures, or more likely eliminates the generality.
The distribution of chlorophyll a is also not surprising. Phytoplankton populations are large in the Rhode River
during the spring and summer seasons . Phytoplankton blooms dominated by dinoflagellates occur frequently
in the Rhode River and are often found following nitrogen inputs during spring runoff or rainfall. We did not
see peaks in chlorophyll a during the spring of 1981, however, probably because it was a drought year and the
usual input of nutrients during spring runoff did not occur. The highest chlorophyll a values (greater than 200
μg chlorophyll a 1-1 attest to the occurrence of such blooms.
The variability of both bacteria and chlorophyll a is related to nutrient concentrations as well as physical
processes. Significant negative correlation coefficients were found for bacteria and chlorophyll with dissolved
inorganic nitrogen components. Nitrogen is likely limiting during most of the summer in the Rhode River  as
it is generally in Chesapeake Bay . The negative correlation probably reflects the rapid uptake of nitrogen
by the plankton community. Positive correlations of phosphorus nutrients with bacteria and chlorophyll reflect
summer peaks of these parameters.
Phytoplankton comprise a significant portion of the microbial community, while bacteria contribute little. If we
use a conversion factor of 250 to estimate total living microbial carbon from the ATP determinations , then
the mean value for total living microbial carbon found in this study is 1.58 mg C 1-1 (range: 0.16-10.78 mg C
1-1). Similarly, a conversion of 30 for chlorophyll a to carbon  yields a mean phytoplankton biomass of 1.26
mg C 1-1 (range: 0.02-7.34 mg C 1-1). Thus, algal biomass constitutes approximately 80% of the total microbial
biomass, and this explains the high positive correlations of chlorophyll a with particulate organic phosphorus
and nitrogen (Table 4), as well as the high positive correlations found with ATP and ETS. A conversion factor
for bacteria, 5.16 × 10-15 g C cell-1, derived from values given in Ferguson and Rublee  and a mean cell
volume estimate of 0.06 μm3 , yields a mean value of 0.06 mg C 1-1 (range: < 0.01-0.29 mg C 1-1). Bacteria
then, contribute only about 4% of the total living microbial carbon. The value for bacteria is similar to that
found by Bell and Albright  in the Strait of Georgia, Canada, but is only about half that reported by Ferguson
and Rublee  for North Carolina coastal waters. Note, however, that the conversion factor used in this study
is conservative; most authors use factors about twice as large [e.g., 3, 9].
Production of the bacteria can be estimated from the thymide incorporation data. To do so we have followed the
protocol of Fuhrman and Azam [13, 14], and used the complete range of conversion factors they suggested,
although recent studies suggest that the high end of the range may be correct [5, 22]. Even under this
conservative approach the productivity of bacteria is often high, with mean values for bacterial production of
0.05-2.09 × 109 cells 1-1 h-1. These values compare favorably with those reported by Fuhrman and Azam 
and Newell and Fallon  for coastal areas. They also span nearly the same range reported by Meyer-Reil 
for the Kiel Bight, although he used a technique of microscopic observation of biomass production in incubated
samples. Finally, the values for production are similar to those reported by Ducklow  for the York River
estuary, Virginia, which is a subestuary of Chesapeake Bay as is the Rhode River. Ducklow also calculated
specific growth rates for bacteria by division of the production estimate by the corresponding standing crop. In
the Rhode River, mean values for such specific growth rates range from 0.26-1.69 d-1 over the range of
conversion factors provided by Fuhrman and Azam  and these are similar to the range of 0.2-1.1 d-1 found
With some estimate of the specific growth rate, speculative determinations of fluxes through the bacterial
assemblage can be attempted. For example, the assimilation efficiency for bacteria is generally taken to be
about 50% . Under this assumption, the net bacterial production is in the range of 0.020.10 mg C 1-1 day-1
and gross production would be twice that. This result suggests several important interactions within the Rhode
River system. First, the source of carbon for bacterial metabolism may be largely derived from algal production.
Williams  and Newell and Fallon  have drawn this conclusion in their studies. The similar overall
correlation between thymidine incorporation and chlorophyll a as compared to bacterial biomass supports this
contention (Table 5). Further, higher correlations of thymidine incorporation with chlorophyll a at the upstream
stations (Table 5) are also consistent with this idea. These stations had the highest chlorophyll values and
therefore primary production which would in turn influence bacterial activity. Second, since production is not
manifested in consequent large bacterial biomass downstream, then losses must be occurring due to settling,
autolysis, or predation. Rublee et al.  have estimated minor losses from the tidal creek into adjacent marsh
areas, but they are not of sufficient magnitude to account for the large production estimate. Settling directly to
the sediment surface is not likely to account for the production either, as tidal and meterological forces would
serve to maintain cells in suspension, and many free-living bacteria would have low sinking rates. Autolysis and
predation have not been evaluated, but we favor predation as a major pathway for recycling. Protozoan
populations are evident in the Rhode River  and many studies have suggested that these organisms are
important trophic links within estuarine systems [e.g., 25, 36].
An additional important feature of the heterotrophic production is that the majority of activity appears to be
associated with free-living bacteria, and not those attached to the particles. We found most cells to be free of
particles during our study, and the highest percentage of activity was located in the 0.2-1.0 μm size fraction.
These results provide an interesting point of comparison with a number of other studies that have assayed for
bacterial numbers and biomass. Generally, in marine systems, less than 5-10% of the heterotrophic activity
appears to be associated with particles [cf. 4, 21]. Hanson and Wiebe  found that most of the heterotrophic
activity in nearshore waters of Georgia was associated with particles. Bell and Albright  found decreasing
proportions of attached bacteria as they moved down the Fraser River estuary. Kirchman and Mitchell  did
not always find the majority of bacteria associated with particles in New England estuarine systems, but they
did find correlations of numbers of attached bacteria with tidal range and the amount of suspended particulate
Our finding of relatively low thymidine incorporation associated with particles in an estuarine system does not
entirely agree with these earlier results, but it is not totally at odds with them either. In fact, on a few occasions
we did find large numbers of bacteria associated with particles and on those occasions the heterotrophic activity
was also predominantly in the larger size range. For example, during the transect of 6/24/81 at station 8, we
found high concentrations of suspended particulates and observed that about 50% of the bacteria were attached
to particles, in contrast to the general pattern of predominantly free-living forms. The incorporation of
thymidine in this sample was also predominantly in larger size fractions than generally expected (Table 3).
Clearly, bacteria-particle interactions are complex and the factors that govern them remain fruitful areas for
1. American Public Health Association (1976) Standard methods for the examination of water and wastewater.
14th ed. Washington, D.C.
2. Azam F, Fenchel T, Field JG, Gray JS, Meyer-Reil LA, Thingstad F (1983) The ecological role of water-
column microbes in the sea. Mar Ecol Progr Ser 10:257-263
3. Bell CR, Albright LJ (1981) Attached and free-floating bacteria in the Fraser River Estuary, British
Columbia, Canada. Mar Ecol Progr Ser 6:317-327
4. Bell CR, Albright LJ (1982) Attached and free-floating bacteria in a diverse selection of water bodies. Appl
Environ Microbiol 43:1227-1237
5. Bell RT, Ahlgren GM, Ahlgren I (1983) Estimating bacterioplankton production by measuring [3H]
thymidine incorporation in a eutrophic Swedish lake. Appl Environ Microbiol 45:17091721
6. Burney CM, Sieburth J McN (1977) Dissolved carbohydrates in seawater. II. A spectrophotometric procedure
for total carbohydrate analysis and polysaccharide estimation. Mar Chem 5:15-28
7. Correll D (1975) The Rhode River program. In: Estuarine Pollution Control and Assessment. Vol I.
Environmental Protection Agency Symposium. Pensacola, Florida, pp 19-27
8. Dobbs RA, Williams RT (1963) Elimination of chloride interference in the chemical oxygen demand test.
Anal Chem 35:1064-1067
9. Ducklow HW (1982) Chesapeake Bay nutrient and plankton dynamics. I. Bacterial biomass and production
during spring tidal destratification in the York River, Virginia estuary. Limnol Oceanogr 27:651-659
10. Faust MA, Correll DL (1976) Comparison of bacterial and algal utilization of orthophosphate in an
estuarine environment. Mar Biol 34:151-162
11. Faust MA, Correll DL (1977) Autoradiographic study to determine metabolically active phytoplankton and
bacteria in the Rhode River estuary. Mar Biol 41:293-305
12. Ferguson RL, Rublee P (1976) Contribution of bacteria to standing crop of coastal plankton. Limnol
13. Fuhrman JA, Azam F (1980) Bacterioplankton secondary production estimates for coastal waters of British
Columbia, Antarctica, and California. Appl Environ Microbiol 39:10851095
14. Fuhrman JA, Azam F (1982) Thymidine incorporation as a measure of heterotrophic bacterioplankton
production in marine surface waters: evaluation and field results. Mar Biol 66: 109-120
15. Haines EB (1979) Interactions between Georgia salt marshes and coastal waters: a changing paradigm. In:
Livingston RJ (ed) Ecological processes in coastal and marine systems. Plenum Press, New York, pp 35-46
16. Hanson RB, Wiebe WJ (1977) Heterotrophic activity associated with particulate size fractions in a Spartina
alterniflora salt marsh estuary, Sapelo Island, Georgia, USA, and continental shelf waters. Mar Biol 42:321-330
17. Hobbie JE, Daley RJ, Jasper S (1977) Use of nuclepore filters for counting bacteria by fluorescence
microscopy. Appl Environ Microbiol 33:1225-1228
18. Holm-Hansen 0, Booth CR (1966) The measurement of adenosine triphosphate in the ocean and its
ecological significance. Limnol Oceanogr 11:510-519
19. Johnson KM, Sieburth J McN (1977) Dissolved carbohydrates in seawater. I. A precise spectrophotometric
analysis for monosaccharides. Mar Chem 5:1-13
20. King EJ (1932) The colorimetric determination of phosphorus. Biochem J 26:292-297
21. Kirchman D, Mitchell R (1982) Contribution of particle-bound bacteria to total microheterotrophic activity
in five ponds and two marshes. Appl Environ Microbiol 43:200-209
22. Kirchman D, Ducklow H, Mitchell R (1982) Estimates of bacterial growth from changes in uptake rates and
biomass. Appl Environ Microbiol 44:1296-1307
23. Maciolek JA (1962) Limnological organic analyses by quantitative dichromate oxidation. Fish Wild Sery
24. Meyer-Reil L (1977) Bacterial growth rates and biomass production. In: Rheinheimer G (ed) Microbial
ecology of a brackish water environment. Springer-Verlag, Berlin, pp 223-236
25. Newell SY, Fallon RD (1982) Bacterial productivity in the water column and sediments of the Georgia
(USA) coastal zone: estimates via direct counting and parallel measurement of thymidine incorporation. Microb
26. Nixon SW (1981) Remineralization and nutrient cycling in coastal marine ecosystems. In: Neilson BJ,
Bronin LE (eds) Estuaries and nutrients. Humana Press, Inc, pp 111-138
27. North BB (1975) Primary amines in California coastal waters: utilization by phytoplankton. Limnol
28. Owens TG, King FD (1975) The measurement of respiratory electron transport system activity in marine
zooplankton. Mar Biol 30:27-36
29. Palumbo AV (1980) Dynamics of bacterioplankton in the Newport River Estuary. PhD Thesis, North
Carolina State University, Raleigh
30. Rublee PA, Merkel SM, Faust MA (in press) Nutrient flux in the Rhode River: tidal transport of
microorganisms in brackish marshes. Est Coastal Shelf Sci
31. Rublee PA, Merkel SM, Faust MA (1983) Transport of bacteria in the sediments of a temperate marsh. Est
Coastal Shelf Sci 16:501-509
32. Sawyer TK, MacLean SA, Coats W, Hilfiker M, Riordan P, Small EB (1976) Species diversity among
sarcodine protozoa from Rhode River, Maryland following tropical storm Agnes. In: CRC Publ No 54, The
effects of tropical storm Agnes on the Chesapeake Bay estuarine system, pp 531-543
33. Shoaf WT, Lium BW (1976) Improved extraction of chlorophyll a and b from algae using dimethyl
sulfoxide. Limnol Oceanogr 21:926-928
34. Strickland JDH (1960) Measuring the production of marine phytoplankton. Bull Fish Res Bd Canada 122:1-
35. Taft JL, Taylor WR (1976) Phosphorus dynamics in some coastal plain estuaries. In: Wiley M (ed)
Estuarine processes. Academic Press, New York, pp 79-90
36. Williams PJ leB (1981) Incorporation of microheterotrophic processes into the classical paradigm of the
planktonic food web. Kiel Meeresfor Sonderh 5:1-28