Luận án tiến sĩ: Structure and function of microbial communities processing dissolved organic matter in marine environments

39 Figure 2.6 Relative abundance of each bacterial group as a function of the fraction of bacteria in phylogenetic group assimilating A glucose and B EPS ….... In the Delaware Estuary th

STRUCTURE AND FUNCTION OF MICROBIAL COMMUNITIES PROCESSING DISSOLVED ORGANIC MATTER IN MARINE

ENVIRONMENTS

by Hila Elifantz

A dissertation submitted to the Faculty of the University of Delaware in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Marine

Studies

Fall 2006

Copyright 2006 Hila Elifantz All Rights Reserved

UMI Number: 3247689

3247689 2007

UMI Microform Copyright

All rights reserved This microform edition is protected against unauthorized copying under Title 17, United States Code.

ProQuest Information and Learning Company

300 North Zeeb Road P.O Box 1346 Ann Arbor, MI 48106-1346

by ProQuest Information and Learning Company

STRUCTURE AND FUNCTION OF MICROBIAL COMMUNITIES PROCESSING DISSOLVED ORGANIC MATTER IN MARINE

ENVIRONMENTS

by Hila Elifantz

I certify that I have read this dissertation and that in my opinion it meets the academic and professional standard required by the University as a

dissertation for the degree of Doctor of Philosophy

Signed:

David L Kirchman, Ph.D

Professor in charge of dissertation

I certify that I have read this dissertation and that in my opinion it meets the academic and professional standard required by the University as a

dissertation for the degree of Doctor of Philosophy

Signed:

David A Hutchins, Ph.D

Member of dissertation committee

I certify that I have read this dissertation and that in my opinion it meets the academic and professional standard required by the University as a

dissertation for the degree of Doctor of Philosophy

Signed:

Thomas Hanson, Ph.D

Member of dissertation committee

I certify that I have read this dissertation and that in my opinion it meets the academic and professional standard required by the University as a

dissertation for the degree of Doctor of Philosophy

Signed:

Byron C Crump, Ph.D

Member of dissertation committee

ACKNOWLEDGMENTS

I would like to thank my advisor David Kirchman, for guiding me in the marine microbial world and for his patience with my writing My committee; David Hutchins, Thomas Hanson, and Byron Crump for their constructive suggestions regarding my research and my manuscripts along the way I would also like to thank Steve Wilhelm from the University of Tennessee for the chlorophyll data

I would like to thank the members of Kirchman lab Liying Yu , Lisa Waidner, Rex Malmstrom, Paul Jones, Vanessa Michelou, Barbara Campbell, Matthew Cottrell, Tiffany Straza, Glen Christman, Dawn Castle and Katie Preen All were very helpful in collecting samples, lab work, and in discussing ideas for this study and the results that came out of it

I would like to thank Coren Milbury, Robin Varney, and Nathan Campbell for accommodating my samples on the sequencer Also thank to the people in Marsh,

Warner, Targett, Cary, Sharp, and Hanson labs for letting me use various instruments in their labs Thanks also for the CMES stuff, especially Peggy Conlon, Paul Dumigan, Lisa Perelly, Doris Manship, Susan Wedeman,

Thanks for all my friends in CMS for making the time here enjoyable Special thanks to Elif Demir, Karen Pelletreau, Tracy Szela, Rick and Linda Rouf for their support throughout and especially at the end when it seemed that I will never finish Finally, thank you my friends and family in Israel, for your support from far and the patience

TABLE OF CONTENTS

LIST OF TABLES……… vii

LIST OF FIGURES……… viii

ABSTRACT……… x

1 INTRODUCTION……… 1

References……… 8

2 ASSIMILATION OF POLYSACCHARIDES AND GLUCOSE BY MAJOR BACTERIAL GROUPS IN THE DELWARE ESTUARY……… 13

Abstract……… 13

Introduction……… 14

Materials and Methods……… 16

Preparation of 3H-EPS……… 16

Sample collection and preparation……… 17

FISH and microautoradiography analysis……… 17

Results……… 19

Dominant bacterioplankton groups assimilating glucose and

EPS……… 19

Relationships between abundance and DOM assimilation……… 21

The importance of EPS and glucose assimilation to a phylogenetic group……… 22

Cell volumes of active and inactive bacteria……… 23

Discussion……… 24

References……… 28

3 DISSOLVED ORGANIC MATTER ASSIMILATION BY HETEROTROPHIC BACTERIAL GROUPS IN THE WESTERN ARCTIC OCEAN……… 41

Abstract……… 41

Introduction……… 42

Materials and Methods……… 45

Sample collection and uptake of 3H-DOM……… 45

FISH and microautoradiography analysis……… 46

Data analysis ……… 47

Results……… 47

Abundance of bacterial groups……… 47

Fraction of cells assimilating DOM components……… 48

Fraction of phylogenetic groups assimilating DOM components 49 Contribution of phylogenetic groups to DOM uptake………… 50

Relationships between abundance and DOM assimilation……… 52

Discussion……… 53

References……… 58

4 DIVERSITY AND ABUNDANCE OF GLYCOSYL HYDROLASES FAMILY 5 IN THE NORTH ATLANTIC OCEAN……… 70

Abstract……… 70

Introduction……… 71

Materials and Methods……… 74

Sample collection……… 74

DNA extraction……… 75

PCR primers design……… 75

GH5 libraries construction……… 76

GH5 libraries sequencing……… 77

Diversity analysis……… 77

Quantitative PCR……… 78

Additional analyses……… 79

Results……… 80

Degenerate GH5 primer design……… 80

Diversity analysis of GH5 clone libraries ……… 81

Abundance of GH5 in the North Atlantic Ocean……… 84

Discussion……… 85

References……… 93

5 CONCLUSIONS……… 109

References……… 114

LIST OF TABLES

Table 2.1 Averaged cell volumes of EPS assimilating and

non-assimilating bacterial cells……… 33

Table 2.2 Averaged cell volumes of glucose assimilating and

non-assimilating bacterial cells……… 34

Table 3.1 Relative abundance of bacteria and major groups in the Western

Arctic Ocean……… 63 Table 4.1 GH5 clusters as classified by amino acids similarity……… 98 Table 4.2 Accession numbers of sequences used in Figure 4.2……… 99 Table 4.3 Sub-groups of cloned GH5 from the MAB and SAR libraries as

appeared in the GH5 gene tree (Figure 4.4)……… 100

LIST OF FIGURES

Figure 2.1 Percent cell contributing to DOM assimilation verses

percent silver grain area around these cells in each

Figure 2.2 Composition of bacterial communities assimilating A)

glucose and B) EPS in the Delaware Estuary………… 36

Figure 2.3 SAR11 clade abundance and contribution to glucose and

EPS assimilation in the Delaware Estuary……… 37

Figure 2.4 Contribution of phylogenetic groups to A) glucose and B)

EPS assimilation as a function of group abundance…… 38

Figure 2.5 Fraction of bacteria in group assimilating glucose vs the

fraction of bacteria in group assimilating EPS………… 39

Figure 2.6 Relative abundance of each bacterial group as a function of

the fraction of bacteria in phylogenetic group assimilating A) glucose and B) EPS … 40 Figure 3.1 Sampling sites in the Western Arctic Ocean……… 64

Figure 3.2 Fraction of all assimilating cells as a function of exposure

time to photographic emulsion……… 65

Figure 3.3 Fraction of bacteria in group that assimilated DOM A)

Cytophaga-like bacteria; B) Alpha-proteobacteria; C)

Gamma-proteobacteria……… 66

Figure 3.4 A) Average percent assimilation of DOM by bacterial

group as a function of location B) Total number of cells in bacterial groups in summer 2004……… 67

Figure 3.5 The contribution to DOM assimilation by specific bacterial

Figure 4.3 Number of operational taxonomic units as a function of

number of clones at 97% similarity A) Nucleic acids; B)

Figure 4.4 Neighbor-joining GH5-like gene tree……… 104

Figure 4.5 Percent of clones in each GH5 library that were grouped in

the same operational taxonomic unit with sequences from

Figure 4.6 Alignment of the amino acid sequence of CelZ from

Erwinia chrysanthemi and the GH5 consensus from MAB

and SAR libraries……… 106

Figure 4.7 Abundance of GH5 genes and chlorophyll concentration

along a transect in the North Atlantic Ocean……… 107 Figure 4.8 Abundance of GH5 genes and chlorophyll concentration in

a depth profile from the Sargasso Sea……… 108

respiration, dissolved organic matter (DOM) concentrations and fluxes However, none

of these methods reveal which bacterial groups use specific compounds in the DOM pool The aim of this dissertation was to address some of these issues

Microautoradiography combined with fluorescence in situ hybridization FISH) was used to evaluate which bacterial groups assimilate common DOM

(Micro-components, and whether this assimilation can be related to other environmental factors

In the Delaware Estuary the assimilation of glucose and extracellular polymeric

substances (EPS) was dominated by the abundant bacterial groups In the freshwater end

of the estuary Actinobacteria and Betaproteobacteria were the dominant groups, while in the saline part of the estuary Alphaproteobacteria and Cytophaga-like bacteria

contributed the most to DOM uptake Only 35-50% of the assimilation could be

explained by bacterial group abundance In addition, groups that

had more assimilating cells were not necessarily more abundant Therefore, it seems that the bacterial community in the Delaware Estuary might be affected equally by "bottom-up" (DOM availability) and "top-down" (predation and viral lysis) factors

In contrast, the oligotrophic environment of the Western Arctic Ocean displayed a different pattern Most of the assimilation (90-99%) could be explained by abundance in the Arctic Ocean These data suggest that the microbial community in this environment is mostly controlled by DOM availability Perhaps the most interesting observation of this study was the level of activity detected in the Arctic environment Up to 50% of all prokaryotes assimilated amino acids, followed by EPS and proteins Glucose, however, was the least assimilated compound The contribution to assimilation of DOM was different among the bacterial groups The contribution to DOM assimilation by

Cytophaga-like bacteria decreased between the Chukchi Sea shelf and the Canada Basin

In contrast, Alphaproteobacteria contributed the most in the slope region This group also

contributed more to the assimilation of low molecular weight (LMW) DOM, while

Cytophaga-like bacteria contributed more to the assimilation of high molecular weight

(HMW) DOM

The last part of this dissertation dealt with the molecular mechanism of the

degradation of polysaccharides, which is an important part of the labile DOM pool in the marine environment The Carbohydrate-Active enZymes (CAZy) and the Sargasso Sea whole-shotgun databases were searched for the common endoglucanases in the marine environment One sub-group of glycosyl hydrolases, family 5 (GH5), was identified as a potential important enzyme in the marine environment Two GH5 gene-libraries were

constructed from the Mid-Atlantic Bight and the Sargasso Sea The two libraries were different from each other The sequences in these libraries were also different from sequences known from cultured bacteria Translation of the amplified fragment indicated that the important residues for activity are present, and therefore these are potentially active endoglucanases GH5 abundance in surface water of the North Atlantic, as

measured with quantitative PCR (Q-PCR), fluctuated between 10 copies to 200 copies per nanogram DNA The abundance of GH5 correlated to chlorophyll concentrations in the eastern part of the sampled region and in one depth profile

The current study added to the growing information regarding the composition of bacterial community in aquatic environments and the role of specific bacterial groups in DOM assimilation In particular, this study was the first to unfold the relation between structure and function of the bacterial community in the Arctic Ocean, the only cold environment studied in that aspect to date The molecular study of GH5 revealed the potential of the community for polysaccharides degradation, however, more need to be done to broaden our understanding of the mineralization of these compounds in the marine environment

microzooplankon (Sherr et al 2002) Therefore, understanding the fate of DOM cycled through the microbial loop is essential for the future assessment of carbon cycling and sequestration in a changing environment

One of the major identified components of the biologically fixed carbon in the marine environment is polysaccharides (Benner et al 1992; Skoog and Benner 1997)

About 20% of the high molecular weight dissolved organic matter (HMW DOM) is composed of polysaccharides, which are produced by organisms from all kingdoms Perhaps the most abundant polysaccharide in the marine environment is chitin, which is the second-most abundant biopolymer on earth (Gooday 1990) and composes half of the HMW dissolved organic nitrogen in ocean surface water

(Aluwihare et al 2005) While chitin is produced mostly by animals (Gooday 1990) and some diatoms (Hoagland et al 1993; Smucker 1991), several structural

polysaccharides, such as agar, carrageen, laminarin, fucoidan, mannan, and xylan, are produced by macro-algae (Giordano et al 2006) A large fraction of the

polysaccharides, however, cannot be identified by a specific structure But, the monosaccharide composition of these polysaccharides suggests that they originate from plants, either terrestrial or marine (Biersmith and Benner 1998; Opsahl and Benner 1999) The use of polysaccharides as a carbon and energy source for the heterotrophic marine bacteria is suggested by their decreasing concentrations with depth (Amon and Benner 1994)

Phytoplankton and bacteria in the marine environment produce extracellular polymeric substances (EPS) which are composed mostly of polysaccharides

(Hoagland et al 1993; Staats et al 1999) While the monosaccharide composition of EPS has been characterized (Taylor et al 1999; Underwood et al 2004), the structure

of polysaccharides in EPS remains unknown A few roles of these polymers have been suggested and include creation of a microenvironment for nutrient capture, protection from biofouling and extreme conditions, and as a carbon sink for

photosynthesis (Wotton 2004) The composition and production of EPS is therefore influenced by various factors, such as growth phase, light-dark periodicity, nutrient

limitation, and salinity changes (Myklestad 2000) The role of EPS in carbon cycling

is interesting because it can be massively produced by phytoplankton blooms (Fajon

et al 1999) The fate of this EPS can be either degradation by the heterotrophic

community or aggregation with microbial cells to form particles and marine snow, which eventually sinks to the ocean interior as part of the biological pump (Engel et

al 2004; Simon et al 2002) It is yet unclear what fraction of the EPS is degraded by the prokaryotic community and what portion of it escapes this degradation The identity of these degrading prokaryotes is also unknown

The interactions between primary and secondary production are important to determine effects of bacteria on DOM in time and space These interactions can be either coupled or decoupled and have been documented in various regions of the oceans and marine ecosystems Coupling is defined as a significant correlation

between dissolved primary production and bacterial production If bacterial carbon demand is much higher than dissolved primary production, then bacteria must have others sources of C However, if dissolved primary production can supply the

bacterial carbon demand, phytoplankton and bacteria are considered to be coupled (Moran et al 2002) In the low productivity water of the Southern Ocean,

heterotrophic bacteria are tightly coupled to phytoplankton through DOC (Moran et

al 2001) In contrast, the coastal regions of Antarctica experience a lag of about a month between phytoplankton bloom and the maximum of bacterial activity,

indicating decoupling due to lack of nutrients (Carlson et al 1998; Moran et al 2001) Similarly, an uncoupling between prokaryotic productivity or respiration and primary production occurs in temperate oligotrophic environments and upwelling zones, leading to overall net heterotrophy of the system (Aristegui and Harrison 2002;

Hoppe et al 2002; Schauer et al 2003) While in the coastal regions of Antarctica the result of the uncoupling is accumulation of DOC, in temperate environments and upwelling zones this uncoupling indicates the use of carbon sources that were

produced elsewhere

About half of the DOM released by primary producers is processed by

heterotrophic bacteria (Azam 1998) The consumption rates of DOM can be evaluated indirectly by measurements of respiration, changes in DOC concentration, bacterial growth rates, and bacterial uptake of radioactive compounds (Williams 2000) The bacterial assimilation of radioactive compounds can be used to estimate fluxes of labile compounds (hours-days), while changes in DOC concentrations assess the flux

of the semilabile fraction (weeks-months) As most freshly produced DOM is labile (Amon and Benner 1994), it is rapidly consumed (Kirchman 2004) The semilabile DOC may be an important source of carbon and other elements for bacterial growth when labile DOC is not available The identity of the bacteria involved in DOC processes was unknown until recently, mostly due to methodological limitations The methods described above treat the bacteria as one unit ("black box") instead of

individuals and cannot identify key organisms in these processes (Azam 1998;

Fasham et al 1999; Gonzalez et al 2003)

The rapid development of molecular techniques at the end of the 1980's – early 1990's increased tremendously our knowledge regarding the bacterial

community composition in aquatic environments The diversity of 16S rRNA genes suggests that the bacterial community is more diverse that it was thought to be based

on data from cultured bacteria (Giovannoni and Rappe 2000; Schmidt et al 1991) The most abundant bacterial group found in clone libraries of 16S rRNA genes in

marine environments is the Alphaproteobacteria, with the SAR11 and Roseobacter

clusters being the most ubiquitous (Giovannoni and Sting 2005) Other groups

represented in 16S rRNA clone libraries and metagenomics libraries include

Gammaproteobacteria, Actinobacteria, picophytoplankton, and three other groups

that are dominant in the aphotic zone (Giovannoni and Rappe 2000) Some groups

such as Cytophaga-like bacteria are underrepresented in clone libraries (Cottrell and

Kirchman 2000a), probably due to PCR biases In a recent analysis of the

accumulated 16S rRNA sequences in the database, it was stressed that some taxa are strictly endemic to one environment, while others are cosmopolitan (Pommier et al 2005) This observation is particularly important when the abundance and role of specific taxa in various environments are to be determined

The accumulated information about 16S rRNA genes allowed the

development of molecular probes to quantify specific phylogenetic bacterial or archaeal groups using fluorescence microscopy In contrast to gene libraries,

fluorescence in situ hybridization (FISH) can be used to determine the abundance in multiple samples relatively quickly (Amann et al 1995; Glöckner et al 1999) In

general, Betaproteobacteria and Actinobacteria are the dominant bacterial groups in

freshwater, either lakes or at the freshwater end of estuaries (Bouvier and Del Giorgio 2002; Glöckner et al 1999; Kirchman et al 2005) With the increase of salinity

Alphaproteobacteria become more abundant In contrast, Cytophaga-like bacteria are

abundant throughout the salinity gradient However, the dominance of these groups can vary with environment In the Antarctic Ocean and coastal Pacific Ocean,

Cytophaga-like bacteria are the most abundant (Cottrell and Kirchman 2000a;

Glöckner et al 1999) In other marine environments and a few lakes this group and

Alphaproteobacteria are equally abundant, while Gammaproteobacteria are the least

abundant (Glöckner et al 1999)

The feasibility of the FISH technique and the growing sequence database led

to the development of yet another useful method for exploring the role of

heterotrophic bacteria in DOM cycling By using a combination of autoradiography and FISH, it became possible to track which bacterial groups take up specific DOM compounds (Lee et al 1999; Ouverney and Fuhrman 1999) Although

autoradiography was not new, and some studies used it to detect bacterial cells

assimilating leucine or thymidine (Smith and Del Giorgio 2003), the use of FISH opened the method for more indepth inquiries Various names were given to this method including MAR-FISH (Lee et al 1999), STAR-FISH (Ouverney and Fuhrman 1999), and Micro-FISH (Cottrell and Kirchman 2000b) Since its establishment, Micro-FISH has been used in various studies to determine the role of bacteria and archaea in the assimilation of DOM components Some general observations can be made from the data collected so far

Archaea actively assimilate various DOM components, including amino acids, glucose, protein and polysaccharides (Kirchman et al In press; Ouverney and

Fuhrman 2000) In contrast, Alphaproteobacteria, and in particular the SAR11

cluster, assimilate more LMW than HMW DOM In the Delaware Estuary this group

assimilates more amino acids and N-acetylglucosamine than the HMW compounds

these materials are derived from (Cottrell and Kirchman 2000b) In various

environments SAR11 dominates the assimilation of dimethylsulfoniopropionate (Malmstrom et al 2004a; Malmstrom et al 2004b; Vila et al 2004) and amino acids

(Malmstrom et al 2005) Cytophaga-like bacteria seem to be important in the

assimilation of HMW DOM (Cottrell and Kirchman 2000b) However, not enough information regarding this group is available to draw general conclusions about its role in the aquatic environment Even less is known about other bacterial groups

The main subject of this study is the processing of polysaccharides in the marine environment Although the importance of polysaccharides in the DOM pool is well established (see above), little is known about the bacterial groups that use these compounds and what processes are involved Two main approaches were used here to gain more knowledge on this subject The isolation of radiolabeled EPS from a

diatom and its use in Micro-FISH allowed the identification of the groups that use this compound However, since polysaccharides are not the only component of the DOM pool, other compounds such as amino acids, glucose and protein were used as well Two different environments were examined during this study: the eutrophic temperate Delaware Estuary (Chapter 2) and the oligotrophic cold western Arctic Ocean

(Chapter 3) Eutrophic environments are well studied; however, these systems and the processes occurring within them are not completely understood The oligotrophic Arctic Ocean is an interesting environment due to its potential role in global warming (Kerr 1999) The second approach used here to explore polysaccharides processes included a molecular study of the abundance and diversity of glycosyl hydrolases in the North Atlantic Ocean (Chapter 4) A search of available sequence resources was done to establish the important enzyme groups in this environment Subsequently, two clone libraries from Delaware coastal waters and Sargasso Sea were constructed and compared Finally, quantitative PCR (Q-PCR) was employed to quantify the amount

of these genes in the North Atlantic Ocean A general conclusion chapter, which

integrates the results from the different chapters as well as future research

possibilities, closes this dissertation

References

ALUWIHARE,L.I., D.J.REPETA, S.PANTOJA, and C.G.JOHNSON 2005 Two

chemically distinct pools of organic nitrogen accumulate in the ocean Science

308: 1007 - 1010

AMANN,R.I., W.LUDWIG, and K.H.SCHLEIFER 1995 Phylogenetic identification

and in situ detection of individual microbial cells without cultivation

Microbiology Reviews 59: 143-169

AMON,R.M.W., and R.BENNER 1994 Rapid cycling of high-molecular-weight

dissolved organic matter in the ocean Nature 369: 549-552

community respiration in the ocean: implications for regional carbon studies

Aquat Microb Ecol 29: 199-209

AZAM,F 1998 Microbial control of oceanic carbon flux: The plot thickens Science

280: 694-696

BENNER,R., J.D.PAKULSKI, M.MCCARTHY, J.I.HEDGES, and P.G.HATCHER 1992

Bulk chemical characteristics of dissolved organic-matter in the ocean

Science 255: 1561-1564

produced dissolved organic matter Mar Chem 63: 131-144

BOUVIER,T.C., and P.A. DEL GIORGIO 2002 Compositional changes in free-living

bacterial communities along a salinity gradient in two temperate estuaries

Limnol Oceanogr 47: 453-470

and C A Carlson [eds.], Biogeochemistry of Marine Dissolved Organic Matter Elsevier Science

carbon partitioning during spring phytoplankton blooms in the Ross Sea

polynya and the Sargasso Sea Limnol Oceanogr 43: 375-386

COTTRELL,M.T., and D.L.KIRCHMAN 2000a Community composition of marine

bacterioplankton determined by 16S rRNA gene clone libraries and

fluorescence in situ hybridization Appl Environ Microbiol 66: 5116-5122

- 2000b Natural assemblages of marine proteobacteria and members of the

Cytophaga-Flavobacter cluster consuming low- and high- molecular-weight

dissolved organic matter Appl Environ Microbiol 66: 1692-1697

export and the biological pump Oceanography 14: 50-58

ENGEL,A., S.THOMS, U.REIBESELL, E.ROCHELLE-NEWALL, and I.ZONDERVAN

2004 Polysaccharide aggregation as a potential sink of marine dissolved

organic carbon Nature 428: 929-932

FAJON,C and others 1999 The accumulation and release of polysaccharides by

planktonic cells and the subsequent bacterial response during a controlled

experiment FEMS Microbiol Ecol 29: 351-363

FASHAM,M.J.R., P.W.BOYD, and G.SAVIDGE 1999 Modeling the relative

contributions of autotrophs and heterotrophs to carbon flow at a Lagrangian JGOFS station in the Northeast Atlantic: The importance of DOC Limnol

Oceanogr 44: 80-94

FIELD,C.B., M.J.BEHRENFELD, J.T.RANDERSON, and P.G.FALKOWSKI 1998

Primary production of the biosphere: Integration terrestrial and oceanic

components Science 291: 237-240

GIORDANO,A., G.ANDREOTTI, A.TRAMICE, and A.TRINCONE 2006 Marine glycosyl

hydrolases in the hydrolysis and synthesis of oligosaccharides Biotechnology

Journal 1: 511-530

GIOVANNONI,S.J., and M.S.RAPPE 2000 Evolution, diversity, and molecular

ecology of marine prokaryotes, p 47-84 In D L Kirchman [ed.], Microbial

Ecology of the Oceans Wiley-Liss, Inc

GIOVANNONI,S.J., and U.STING 2005 Molecular diversity and ecology of microbial

plankton Nature 437: 343-348

GL Ö CKNER,F.O., B.M.FUCHS, and R.AMANN 1999 Bacterioplankton compositions

of lakes and oceans: a first comparison based on fluorescence in situ

hybridization Appl Environ Microbiol 65: 3721-3726

GONZALEZ,N., R.ANADON, and L.VIESCA 2003 Carbon flux through the microbial

community in a temperate sea during summer: role of bacterial metabolism

Aquat Microb Ecol 33: 117-126

GOODAY,G.W 1990 The ecology of chitin degradation Adv Microb Ecol 1:

387-430

organic carbon Global Biogeochemistry Cycles 12: 443-453

HOAGLAND,K.D., J.R.ROSOWSKI, M.R.GRETZ, and S.C.ROEMER 1993 Diatom

extracellular polymeric substances - Function, fine-structure, chemistry,and

physiology J Phycol 29: 537-566

HOPPE,H.G., K.GOCKE, R.KOPPE, and C.BEGLER 2002 Bacterial growth and

primary production along a north-south transect of the Atlantic Ocean Nature

416: 168-171

KERR,R.A 1999 Will the Arctic Ocean Lose All Its Ice? Science 286: 1828

prokaryotes in the oceans, p 31-63 In M Follows and T Oguz [eds.], The

ocean carbon cycle and climate Kluwer Academic Publishers

KIRCHMAN,D.L., A.I.DITTEL, R.R.MALMSTROM, and M.T.COTTRELL 2005

Biogeography of major bacterial groups in the Delaware Estuary Limnol

Oceanogr 50: 1697-1706

KIRCHMAN,D.L., H.ELIFANTZ, A.I.DITTEL, R.R.MALMSTROM, and M.T

COTTRELL In press Standing stocks and activity of archaea and bacteria in the western Arctic Ocean Limnol Oceanogr

LEE,N and others 1999 Combination of fluorescent in situ hybridization and

microautoradiography -a new tool for structure-function analyses in

microbial ecology Appl Environ Microbiol 65: 1289-1297

MALMSTROM,R.R., M.T.COTTRELL, H.ELIFANTZ, and D.L.KIRCHMAN 2005

Biomass production and dissolved organic matter assimilation by SAR11

bacteria in the Northwest Atlantic Ocean Appl Environ Microbiol 71:

2979-2986

MALMSTROM,R.R., R.P.KIENE, M.T.COTTRELL, and D.L.KIRCHMAN 2004a

Contribution of SAR11 bacteria to dissolved dimethylsulfoniopropionate and

amino acid uptake in the North Atlantic Ocean Appl Environ Microbiol 70:

4129-4135

MALMSTROM,R.R., R.P.KIENE, and D.L.KIRCHMAN 2004b Identification and

enumeration of bacteria assimilating dimethylsulfoniopropionate (DMSP) in

the North Atlantic and Gulf of Mexico Limnol Oceanogr 49: 597-606

MORAN,X.A.G., M.ESTRADA, J.M.GASOL, and C.PEDROS-ALIO 2002 Dissolved

primary production and the strength of phytoplankton-bacterioplankton

coupling in contrasting marine regions Microb Ecol 44: 217-223

MORAN,X.A.G., J.M.GASOL, C.PEDROS-ALIO, and M.ESTRADA 2001 Dissolved

and particulate primary production and bacterial production in offshore

Antarctic waters during austral summer: coupled or uncoupled? Mar Ecol

Prog Ser 222: 25-39

In P J Wangersky [ed.], The handbook of environmental chemistry Springer

OPSAHL,S., and R.BENNER 1999 Characterization of carbohydrates during early

diagenesis of five vascular plant tissues Org Geochem 30: 83-94

rRNA probe technique for determination of radioisotope uptake by specific

microbial cell types in situ Appl Environ Microbiol 65: 1746-1752

- 2000 Marine planktonic archaea take up amino acids Appl Environ Microbiol

66: 4829-4833

POMMIER,T., J.PINHASSI, and A.HAGSTROM 2005 Biogeographic analysis of

ribosomal RNA clusters from marine bacterioplankton Aquat Microb Ecol

41: 79-89

SCHAUER,M., V.BALAGUE, C.PEDROS-ALIO, and R.MASSANA 2003 Seasonal

changes in the taxonomic composition of bacterioplankton in a coastal

oligotrophic system Aquat Microb Ecol 31: 163-174

SCHMIDT,T.M., E.F.DELONG, and N.R.PACE 1991 Analysis of a marine

picoplankton community by 16s ribosomal- RNA gene cloning and

sequencing J Bacteriol 173: 4371-4378

SHERR,E.B., B.F.SHERR, and P.G.VERITY 2002 Distribution and relation of total

bacteria, active bacteria, bacterivory, and volume of organic detritus in

Atlantic continental shelf waters off Cape Hatteras NC, USA Deep-Sea Res

Part II-Top Stud Oceanogr 49: 4571-4585

SIMON,M., H.-P.GROSSART, B.SCHWEITZER, and H.PLOUG 2002 Microbial ecology

of organic aggregates in aquatic ecosystems Aquat Microb Ecol 28:

175-211

SKOOG,A., and R.BENNER 1997 Aldoses in various size fractions of marine organic

matter: Implications for carbon cycling Limnol Oceanogr 42: 1803-1813

SMITH,E.M., and P.A. DEL GIORGIO 2003 Low fractions of active bacteria in

natural aquatic communities? Aquat Microb Ecol 31: 203-208

Ecology 19: 357-369

STAATS,N., B.DE WINDER, L.J.STAL, and L.R.MUR 1999 Isolation and

characterization of extracellular polysaccharides from the epipelic diatoms

Cylindrotheca closterium and Navicula salinarum Eur J Phycol 34: 161-169

TAYLOR,I.S., D.M.PATERSON, and A.MEHLERT 1999 The quantitative variability

and monosaccharide composition of sediment carbohydrates associated with

intertidal diatom assemblages Biogeochemistry 45: 303 - 327

UNDERWOOD,G.J.C., M.BOULCOTT, C.A.RAINES, and K.WALDRON 2004

Environmental effects on exopolymer production by marine benthic diatoms:

dynamics, changes in composition, and pathways of production J Phycol 40:

293-304

VILA,M and others 2004 Use of microautoradiography combined with fluorescence

in situ hybridization to determine dimethylsulfoniopropionate incorporation

by marine bacterioplankton taxa Appl Environ Microbiol 70: 4648-4657

WILLIAMS,P.J. L.B 2000 Heterotrophic bacteria and the dynamics of dissolved

organic material In D L Kirchman [ed.], Microbial Ecology of the Oceans

Wiley-Liss

systems Scientia Marina 68: 13-21

Chapter 2

ASSIMILATION OF POLYSACCHARIDES AND GLUCOSE BY MAJOR

BACTERIAL GROUPS IN THE DELWARE ESTUARY

Abstract

The contribution of major bacterial groups to the assimilation of extracellular polymeric substances (EPS) and glucose in the Delaware Estuary was assessed using microautoradiography and fluorescence in situ hybridization (Micro-FISH) Bacterial groups contributed to EPS and glucose assimilation in part according to their

distribution in the estuary Abundance of the phylogenetic groups explained 35% and

55% of the variation in EPS and glucose assimilation, respectively Actinobacteria contributed 70 % to glucose assimilation in freshwater, while Alphaproteobacteria

assimilated 60% of this compound in saline water In contrast, a broad range of

bacterial groups dominated the assimilation of EPS Actinobacteria and

Betaproteobacteria contributed the most in the freshwater section whereas

Cytophaga-like bacteria, Alphaproteobacteria and Gammaproteobacteria participated

in EPS assimilation in the lower part of the estuary In addition, we examined the fraction of bacteria in each group assimilating glucose or EPS Overall, the fraction

of bacteria in all groups that assimilated glucose was higher than the fraction that assimilated EPS (15-30% versus 5-20%, respectively) We found no correlation between relative abundance of a group in the estuary and the fraction of bacteria actively assimilating glucose or EPS; the more active groups were often less

abundant Our results imply that the bacterial community in the Delaware Estuary is not controlled solely by "bottom-up" factors such as dissolved organic matter

Introduction

Carbohydrates are a major component of the dissolved organic matter (DOM)

in marine environments and account for about 20% of dissolved organic carbon, with polysaccharides comprising the bulk of these compounds (Benner et al 1992)

Concentrations of polysaccharides decrease with depth, suggesting that these

compounds are labile (Pakulski and Benner 1994) Concentrations of

monosaccharides like glucose are generally very low (<50 nM) (Borch and

Kirchmann 1997; Skoog et al 1999), but fluxes can be large enough to support more than 30% of bacterial growth in the surface ocean (Carlson 2002) Carbohydrates are produced by active or passive release by phytoplankton (Aluwihare and Repeta 1999; Biersmith and Benner 1998) and other organisms (Bartlett et al 1988) Regimes with high nutrient concentrations, such as upwelling zones and coastal areas, have high production and elevated concentrations of polysaccharides and other extracellular polymeric substances (EPS) (Underwood et al 2004)

Previous studies have examined the uptake of glucose and EPS by the total microbial assemblage (Janse et al 2000; Malmstrom et al 2005), but little is known about the uptake of these compounds by specific bacterial groups In the Atlantic

Ocean, the Alphaproteobacterial group SAR11 accounts for about 50% of glucose

assimilation (Malmstrom et al 2005) The SAR11 clade and other

Alphaproteobacteria are also important in the assimilation of other low molecular

weight (LMW) compounds such as amino acids and N-acetylglucosamine in

Delaware coastal waters (Cottrell and Kirchman 2000), and

dimethylsulfoniopropionate (DMSP) in the Gulf of Mexico, Northwest

Mediterranean, the coastal North Atlantic Ocean, and the Sargasso Sea (Malmstrom

et al 2004a; Malmstrom et al 2004b; Vila et al 2004) Uptake of EPS by specific bacterial groups has not been examined, but a few studies have considered other high

molecular weight (HMW) compounds Cytophaga-like bacteria appear to dominate protein and chitin use in Delaware coastal waters, whereas Alphaproteobacteria were

less important in uptake of these compounds (Cottrell and Kirchman 2000) Likewise, SAR11 bacteria accounted for much less of protein assimilation than of glucose assimilation in the North Atlantic Ocean (Malmstrom et al 2005)

Single cell analyses of leucine and thymidine assimilation have provided some insights into the contribution of specific phylogenetic groups to bacterial production and thus to total DOM uptake Cottrell and Kirchman (Cottrell and Kirchman 2003) found that 50% of the variation in the assimilation of both leucine and thymidine by a bacterial group was explained by its abundance and that the contribution of the

various phylogenetic groups to bacterial production followed the biogeography of these groups in the Delaware Estuary Data from microautoradiography fluorescence

in situ hybridization (Micro-FISH) can also be used to examine the fraction of cells within each phylogenetic group assimilating leucine and thymidine (Cottrell and Kirchman 2003) In the Delaware Estuary, there is no correlation between this

fraction and the relative abundance of bacterial groups (except for

Betaproteobacteria), suggesting that these groups are controlled by factors other than

bottom-up ones The relationship between abundance and assimilation of other organic compounds is unclear

The goal of this study was to identify the main phylogenetic groups that participate in EPS and glucose assimilation in the Delaware Estuary Polysaccharides are the main carbohydrate in the estuary, as glucose and other monosaccharides cannot be detected (<5 nM) (Kirchman and Borch 2003), most likely because they are rapidly consumed by microbes Bacterial community structure changes along the

salinity gradient of the estuary, with Betaproteobacteria and Actinobacteria abundant

in the freshwater section while Alphaproteobacteria dominate the lower part of the

estuary (Cottrell and Kirchman 2003; Kirchman et al 2005) I hypothesized that the most abundant groups in each location dominate uptake of EPS and glucose We found that abundance only partially explained relative uptake of these compounds, suggesting that bacterial communities are controlled by more than just bottom-up

factors, such as DOM concentrations and composition

Materials and Methods

Preparation of 3 H-EPS: EPS was prepared using an axenic culture of the

heterotrophic diatom Nitzschia leucosigma (CCMP 2197) grown on 3H-glucose as the

sole carbon source The diatom culture was grown on 10 µM 3

H-glucose (33 Ci/mmol, Amersham) overnight in the dark in filter-sterilized (0.2 µm pore-size polycarbonate) and autoclaved Sargasso seawater with no additional nutrients Cells were separated from the culture medium by centrifugation at 2500 X g (20˚C, 20 min) and the supernatant containing the EPS was transferred to a clean tube The soluble EPS was separated from the culture medium by precipitation with ethanol (70% final concentration) overnight at -20°C (De Brouwer and Stal 2002) The precipitated EPS

was resuspended in 70% ethanol (4˚C) and precipitated again under the same

conditions This step, which was essential to remove residual monosaccharides, was repeated three times The EPS was resuspended in 3% NaCl, radio-assayed, and tested for carbohydrate concentrations (in glucose equivalents) using the phenol-sulfuric acid assay (Daniels et al 1994)

Sample collection and preparation: Surface water was collected from four locations

along the Delaware Estuary in July 2003 (salinities 0, 13, 21, and 26 PSU) and June

2004 (28 PSU) Water samples (30 ml) were incubated with 2 nM 3H-glucose (33 Ci/mmol, Amersham) for 2 h or with 1.5 µM 3

H-EPS (concentration in glucose equivalents) for 14 h Both treatments were incubated at the in situ temperature in the dark Paraformaldehyde (PFA, 2% final concentration) was added to killed controls

15 min prior to the addition of the 3H-compounds At the end of the incubation, samples were fixed with PFA (2% final concentration), and all samples were stored at 4˚C for 24 h Samples were then filtered onto 0.2-µm-pore-size polycarbonate filters, which were kept in -20˚C until analysis

FISH and microautoradiography analysis: FISH analysis was done using the

following Cy3-labeled probes: Eub338 for Eubacteria (Amann et al 1990), Alf968 for

Alphaproteobacteria (Gloökner et al 1999), Bet42a and Gam42a for

Betaproteobacteria and Gammaproteobacteria, respectively (Manz et al 1992),

CF319a for Cytophaga-like bacteria (Manz et al 1996), HGC96a for Actinobacteria

(Roller et al 1994), and a suite of four probes for SAR11 bacteria (Morris et al

2002) Unlabeled competitors probes were used for Betaproteobacteria and

Gammaproteobacteria (Manz et al 1992) In addition, a negative probe was used to

determine nonspecific binding (Karner and Fuhrman 1997)

Following FISH analysis, samples were subjected to microautoradiography as described by Cottrell and Kirchman (Cottrell and Kirchman 2003) Glucose samples were exposed for 12-24 h, while EPS samples were exposed for 3-6 d These

exposure times were set so that the percent of total cells associated with silver grains was in the range of 15-20% This range was chosen because that was the maximum percent of cells assimilating EPS that could be detected At the end of the exposure time, the slides were developed as described previously (Cottrell and Kirchman 2003) The samples were mounted with a mixture of 4:1 Citifluor (Ted Pella) and Vectashield (Vector Labs) containing 0.5 ng/µl of 4'-6'-diamidino-2-phenylindole (DAPI) stain, and covered with coverslips Slides were stored at -20˚C until

microscopic analysis

Total numbers of cells (DAPI stained), cells affiliated with a specific bacterial group (Cy3-labeled) and cells that assimilated the radiolabled compound (with silver grains developed during microautoradiography) were counted using a semi-automatic microscope and image analysis as described previously (Cottrell and Kirchman 2003) Data were collected from 30 fields of view and the number of cells counted is

indicated in Tables 1 and 2 The percent of total silver grain area was calculated as follows The total silver grain area around the probe-positive cells was summed and divided by the sum of total silver grain area associated with DAPI-stained cells Cell volumes were calculated using the algorithm described by Sieracki et al (Sieracki et

al 1989)

Data expressed as percentage were arcsin-transformed before analysis Model

II regressions were performed to examine the relationship between abundance and silver grain area Cell volumes were log-transformed before analysis

Results

We examined the contribution of various phylogenetic groups to EPS and glucose assimilation along the Delaware Estuary This contribution can be estimated using either the relative number of substrate-assimilating cells or the total silver grain area associated with a bacterial group In our study, these two parameters varied similarly throughout the estuary for both glucose and EPS uptake (Fig 2.1) Although estimates of assimilation as expressed by silver grain area tended to be higher than estimates assessed by cell abundance (Fig.2.1), the slope between relative cell

abundance and silver grain area was not different from one (slope of 1.07 ± 0.05, p>0.05) Therefore, we present the data using the percent of total silver grain area throughout this paper

Dominant bacterioplankton groups assimilating glucose and EPS

The assimilation of glucose was dominated by a few groups that changed

along the salinity gradient of the estuary Actinobacteria accounted for nearly all of

glucose assimilation (70%) in the freshwater (Fig 2.2A) The contribution of this group decreased dramatically as salinity increased The opposite trend was observed

for Alphaproteobacteria While this group contributed only 20% to glucose

assimilation in the freshwater, Alphaproteobacteria accounted for 60% of

assimilation in the highest salinity The rest of the phylogenetic groups examined in

this study assimilated less glucose Betaproteobacteria and Gammaproteobacteria, and Cytophaga-like bacteria accounted for 15-20% of glucose assimilation,

depending on location

While a single group dominated glucose assimilation in the freshwater and saline ends of the estuary, more than one group assimilated EPS at each location

Both Betaproteobacteria and Actinobacteria accounted for 30% of EPS assimilation

in the freshwater and were the dominant EPS users in this part of the estuary (Fig 2.2B) Assimilation by both phylogenetic groups decreased to 5-20% as salinity increased In the lower part of the estuary (21-28 PSU), different groups dominated

EPS assimilation In 21 PSU water, Alphaproteobacteria and Gammaproteobacteria each accounted for 20% of total silver grain area Cytophaga-like bacteria were the primary EPS users in 26 PSU water, followed by Alphaproteobacteria

Alphaproteobacteria were the main EPS consumers in the highest salinity, comprising

55% of total silver grain area (Fig 2.2B) Unlike glucose assimilation, in which the

Alphaproteobacterial contribution increased gradually along the estuary, this group

used EPS quite similarly in most parts of the estuary (20-25% of total assimilation)

In addition to broad phylogenetic groups, we also examined the contribution

of the SAR11 clade, a sub-group of the Alphaproteobacteria (Giovannoni and Rappe

2000), to assimilation of glucose and EPS The SAR11 clade was abundant (20-33%

of all prokaryotes) in the saline part of the estuary (Fig 3) but was near detection limits in freshwater (Kirchman et al 2005) Glucose assimilation by the SAR11 clade was significantly higher than that of EPS (25% and 15%, respectively) in two of three stations (t-test, p<0.05) However, SAR11 bacteria assimilated these compounds less than expected based on abundance (Fig 2.3)

Relationships between abundance and DOM assimilation

We next examined whether assimilation by a phylogenetic group could be explained by its relative abundance in the community About 55% of the variation in the assimilation of glucose was explained by the abundance of the bacterial groups (Fig 2.4A) A smaller fraction of the variation in EPS assimilation (35%) was

explained by abundance (Fig 2.4B) However, when excluding the data points that were more than 20% off the 1:1 line (21% and 12% of the data points for glucose and EPS uptake, respectively), the slopes between silver grain area and the relative

abundance of each phylogenetic group were not different from one (0.890 ± 0.17 and 0.94 ± 0.36 for EPS and glucose, respectively)

A few bacterial groups assimilated glucose and EPS more than expected based

on their abundance For glucose assimilation, these groups include the

Alphaproteobacteria in the lower part of the estuary (salinity of 21-28 PSU) and Actinobacteria in the freshwater (Fig 2.4A) Although Betaproteobacteria,

Gammaproteobacteria and Cytophaga-like bacteria were abundant in some sites,

these groups used glucose as expected or less than expected according to their

abundance (Fig 2.4A) Alphaproteobacteria assimilated more glucose than expected

from their abundance in three locations (Fig 2.4A), but this group assimilated more

EPS than expected (by 2.5-fold) at only one location (Fig 2.4B) Betaproteobacteria (by 4-fold) and Cytophaga-like bacteria (by 2-fold) also assimilated EPS more than

expected from their abundance in 28 and 26 PSU waters, respectively The bacterial groups that assimilated EPS more than expected in the lower part of the estuary were also the groups contributing the most to EPS assimilation (Fig 2.2B)

The importance of EPS and glucose assimilation to a phylogenetic group

The contribution of a specific phylogenetic group to DOM fluxes does not necessarily reflect the importance of a compound to that group To explore this issue,

we calculated the fraction of bacteria in each group that assimilated glucose or EPS, i.e probe-positive cells with silver grains compared with probe-positive cells in that particular group (with and without silver grains) This is different from the previous sections that present the contribution of each bacterial group to EPS or glucose assimilation, i.e probe-positive cells with silver grains divided by all cells with silver

grains For example, Alphaproteobacteria accounted for 35% of EPS assimilation in

26 PSU water, but only 12% of Alphaproteobacteria used EPS in this region of the

with salinity For example, more Betaproteobacteria assimilated EPS in 28 PSU

water, while more cells of this group used glucose in 21 PSU water

The fraction of bacteria in groups assimilating either compound did not explain the abundance of these groups (r = 0.28 for EPS; r = 0.15 for glucose) (Fig 2.6) Groups that had a high fraction of cells assimilating these compounds were not necessarily abundant Similarly, groups that were abundant had unexpectedly low

fractions of cells assimilating either compound (Fig 2.6) Although Actinobacteria

were more abundant in the freshwater part of the estuary, the fraction of this group assimilating glucose was similar throughout the estuary (Fig 2.6A) However,

relatively more Actinobacteria assimilated EPS in the freshwater section (Fig 2.6B) The abundance of Alphaproteobacteria and SAR11 bacteria assimilating both EPS

and glucose was usually lower than expected based on the fraction of active cells A

higher fraction of Alphaproteobacteria assimilated both compounds only at 28 PSU

Cell volumes of active and inactive bacteria

Cell volumes may change during preparation of samples for the Micro-FISH analysis For that reason we chose to examine the ratio of the volumes for active vs inactive cells We assume that any changes in cell volume are cancelled out in the ratio

The ratios between the volumes of substrate-assimilating and non-assimilating cells indicate that bacteria taking up glucose and EPS were larger than non-

assimilating bacteria (t-test, p<0.05) EPS-assimilating bacteria were larger than

non-assimilating bacteria by 20-44% (Table 2.1) Cytophaga-like bacteria had the highest

ratio of assimilating to non-assimilating cell volumes in the EPS treatment;

assimilating cells were 62% larger than non-assimilating ones This was also the case

for Cytophaga-like bacteria cells assimilating glucose (Table 2.2), in which the assimilating Cytophaga-like bacteria were 57% larger than the non-assimilating cells

affiliated with this group The other phylogenetic groups had smaller differences (range of 10-35%) in the glucose treatment Overall, EPS-assimilating bacteria were larger than glucose-assimilating cells by 10-30%

Discussion

A goal of this study was to determine the contribution of bacterial groups in the Delaware Estuary to glucose and EPS assimilation Relative abundance partially determined the contribution of the major bacterial groups to assimilation of these compounds Group abundance explained 35% and 55% of the variation in EPS and glucose assimilation, respectively Similarly, Cottrell and Kirchman (2003) reported that 50% of bacterial production along the Delaware Estuary is explained by group abundance A few bacterial groups had exceptionally high levels of glucose and EPS assimilation relative to their abundance

Unlike EPS, which was assimilated across a broad range of groups, glucose assimilation was generally dominated by a few major groups In particular,

Alphaproteobacteria dominated glucose assimilation in the saline part of the estuary,

and this group assimilated glucose more than expected based on its relative

abundance while most of the bacterial groups assimilated glucose as expected based

on their abundance These data are consistent with previous studies suggesting that

Alphaproteobacteria are important in the assimilation of LMW compounds such as

amino acids, N-acetylglucosamine (Cottrell and Kirchman 2000), and DMSP

(Malmstrom et al 2004b) The ability to use various LMW DOM compounds perhaps

explains the high abundance of this group Alphaproteobacteria are abundant in

marine environments (Bouvier and Del Giorgio 2002; Glöckner et al 1999; Kirchman

et al 2005), and LMW DOM can support a high fraction of bacterial production (Kirchman 2003)

The SAR11 clade, a sub-group of the Alphaproteobacteria, is abundant in the

oceans (Malmstrom et al 2005; Morris et al 2002) as well as in the Delaware Estuary

(Kirchman et al 2005) Although the SAR11 clade accounts for about 50% of glucose assimilation in the Sargasso Sea (Malmstrom et al 2005), it accounted for only 15-20% of glucose assimilation in the Delaware The SAR11 clade is a very diverse group (Acinas et al 2004; Field et al 1997), and the phylotypes of this group in the Delaware Estuary probably differ from those in the Sargasso Sea The Sargasso Sea is oligotrophic, while the Delaware Estuary is eutrophic This difference probably selects for different phylotypes with diverse metabolisms

In contrast to glucose assimilation, several bacterial groups assimilated EPS Potential reasons for this finding include the fact that polysaccharides and other EPS components may be more important carbon sources than glucose Polysaccharides concentrations are relatively high, whereas glucose is too low to be measured (< 5 nM) in the Delaware (Kirchman and Borch 2003) Second, EPS originating from phytoplankton is a complex mixture of compounds, including polysaccharides,

proteins, lipids and perhaps even some LMW compounds (Underwood et al 2004) In addition, the polysaccharides in EPS are probably diverse and composed of various monosaccharides Therefore, different bacterial groups may use different components

of EPS, preventing a single group from dominating EPS assimilation and allowing a broad spectrum of groups to be involved

One of our initial hypotheses was that Cytophaga-like bacteria would

dominate assimilation of EPS This hypothesis was based on previous findings that cultured representatives of this group degrade polymers such as cellulose (Lynd et al 2002) In addition, uncultured bacteria affiliated with this group contribute to the assimilation of protein and chitin in coastal waters (Cottrell and Kirchman 2000) The

high abundance of Cytophaga-like bacteria on particles (Crump et al 1999; Delong et

al 1993) also suggests that these bacteria might be especially important in the cycling

of HMW DOM However, Cytophaga-like bacteria dominated EPS assimilation only occasionally in the Delaware Estuary Since Cytophaga-like bacteria shared EPS

assimilation with other bacterial groups, this DOM component appears to be an important source of carbon to a variety of bacteria rather than to a specific group

We expected that community composition is controlled in part by bottom-up factors Bottom-up control by DOM is supported by the extent to which there is a correlation between group abundance and its contribution to DOM uptake These correlations indicate that relative DOM uptake explains 30-55% of relative abundance and thus of community composition Bottom-up control also implies that bacterial groups with high fractions of substrate-assimilating cells would be more abundant than groups with lower fractions In contrast, most bacterial groups with high

fractions of substrate-assimilating cells had low abundances in the community

Similarly, the relative abundance of bacterial groups did not follow thymidine-active bacteria (Cottrell and Kirchman 2004) or the growth rates of specific groups in the Delaware Estuary (Yokokawa et al 2004), with one exception The abundance of

Betaproteobacteria followed both thymidine assimilation (Cottrell and Kirchman

2004) and growth rate (Yokokawa et al 2004) in this estuary These data are

consistent with bottom-up control of Betaproteobacteria However, we did not

observe any trends with EPS and glucose assimilation, even for Betaproteobacteria,

suggesting that bacterial community structure is not controlled solely by bottom-up factors If so, then top-down factors such as predation and viral lyses may be the main forces shaping bacterial communities in the Delaware and perhaps other estuaries

Predation and viral lyses can be important top-down factors determining the composition of bacterial communities in aquatic environments (Gonzalez et al 1990; Jurgens and Matz 2002; Simek et al 2003; Thingstad and Lignell 1997)

Susceptibility of bacteria to predation is affected by cell size and indirectly by level of activity (Del Giorgio et al 1996; Sherr et al 2002) This may explain why the more active bacteria in the Delaware Estuary were also less abundant (this study, (Cottrell and Kirchman 2004) Differential grazing (Jurgens and Matz 2002) would also

explain the differences in the size of assimilating and non-assimilating cells found in our study The size of bacteria can also affect viral lysis since viruses tend to attack the largest cells in the community (Murray and Jackson 1992; Weinbauer and Hofle 1998) Bacterial abundance is another factor Thingstad and Lignell (1997) postulated with their "killing the winner" hypothesis that viral lysis is highest for abundant bacterial groups The combination of predation and viral lysis, therefore, affect the abundance, size distribution and diversity of bacterial groups in aquatic communities

Our understanding of the abundance and function of specific bacterial groups

in the aquatic environment has increased over the last decade Overall, it seems that abundant groups contribute the most to the use of different DOM components in

various marine environments, with some important exceptions Alphaproteobacteria

are one of the most abundant bacterial groups in the marine environments and have an important role in the uptake of LMW organic compounds In contrast, several groups appear capable of processing HMW compounds, a diversity that may be a reflection

of the complexity of HMW DOM These data suggests that there is some bottom-up control of the community, but the fraction of substrate-assimilating cells in the groups imply that other factors are involved It is likely that a combination of bottom-up