The Pennsylvania State University
The Graduate School
Department of Civil and Environmental Engineering
ANALYSIS OF MICROBIAL COMMUNITIES AND DESIGN OF BIOREACTORS USED
FOR PERCHLORATE REMEDIATION AND BIOHYDROGEN PRODUCTION
A Thesis in
Environmental Engineering
by
Husen Zhang
© 2005 Husen Zhang
Submitted in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
May 2005 The thesis of Husen Zhang has been reviewed and approved* by the following:
Bruce E. Logan Kappe Professor of Environmental Engineering Thesis Advisor Chair of Committee
Mary Ann Bruns Assistant Professor of Crop and Soil Sciences
John M. Regan Assistant Professor of Environmental Engineering
William D. Burgos Associate Professor of Environmental Engineering
Andrew Scanlon Professor of Civil Engineering Head of the Department of Civil and Environmental Engineering
*Signatures are on file in the Graduate School.
ABSTRACT
A chemolithoautotrophic bacterium (strain HZ) was isolated from biofilm samples in an unsaturated-flow, packed-bed reactor treating perchlorate-contaminated groundwater. Dilution- to-extinction method was used for isolation. Purity was initially examined microscopically, and confirmed by identical intergenic ribosomal RNA spacer sequences from multiple clones. Strain
HZ is a Gram-negative, rod-shaped facultative anaerobe that can use oxygen, perchlorate, chlorate, or nitrate as an electron acceptor and hydrogen gas or acetate as an electron donor.
Growth on hydrogen gas was coupled with complete perchlorate (10 mM) reduction to chloride with a maximum doubling time of 8.9 hours. Autotrophic growth with carbon dioxide as the sole carbon source was confirmed by demonstrating that biomass carbon (100.9%) was derived from
14 CO2. Phylogenetic analysis based upon the 16S rRNA sequence indicated that strain HZ
belongs to the genus Dechloromonas within the β subgroup of the Proteobacteria.
Biofilm samples from a pilot-scale, fixed-bed, perchlorate-reducing reactor were
analyzed for microbial species from inoculated and indigenous populations. The bioreactor was
inoculated with Dechlorosoma sp. strain KJ and fed groundwater containing indigenous
microorganisms. The reactor was flushed weekly to remove accumulated biomass. Perchlorate
was reduced to non-detectable levels (< 4 µg L-1) after 26 days of operation and remained so
during proper reactor operation in a six-month long test. Plastic media in the reactor were
collected from top, middle, and bottom locations. Genomic DNA was extracted from successive
washes of thawed biofilm material for PCR-based community fingerprinting by 16S-23S
ribosomal intergenic spacer analysis (RISA). No DNA sequences closely related to that of strain
KJ were recovered. The most intense bands yielded DNA sequences with high similarities (>
98%) to Dechloromonas spp. Other sequences from RISA fingerprints indicated presence of the
iii low G+C Gram-positive bacteria and the Cytophaga-Flavobacterium-Bacteroides (CFB) group.
Fluorescence in situ hybridization (FISH) was also used to examine biofilms using genus-
specific 16S ribosomal RNA probes. Numbers of bacteria hybridizing to the Dechloromonas
probe were most abundant at the biofilm surface (23% of all cells), and decreased as biofilm
depth increased. The bacteria hybridizing to Dechlorosoma probes constituted less than 1 % of
all cells in the biofilms examined, except in the deepest portions where they represented 3-5%.
Biological hydrogen production was investigated using a laboratory-scale, trickle-bed
biofilm reactor. The reactor was inoculated with Clostridium acetobutylicum and fed mineral
medium with glucose as the sole carbon and energy source. The flowrate was 1.6 mL/min
(0.096 L/h), producing a hydraulic retention time of 2.1 min. Continuous hydrogen production
rates were 14, 20, 25, and 27 mL/h at influent glucose concentrations of 1.0, 3.3, 4.5, and 10.5
g/L, respectively. Gas-phase hydrogen concentrations of 70-79% were obtained at influent
glucose concentrations of 1.0, 3.3, 4.5, and 10.5 g/L. The major fermentation byproducts were
acetate and butyrate. The measured hydrogen yields indicated an overall conversion efficiency
of 15-26% based on a theoretical stoichiometry of 4 moles hydrogen from 1 mole of glucose.
-1 -1 The normalized hydrogen production rate was 676 to 1265 (mL-H2)(g-glucose) (L-reactor) , which is within the range of values reported using continuously stirred tank reactors.
Another H2-producing bioreactor was operated at four hydraulic retention times (HRT)
(10, 5, 2.5 and 1 h) and four glucose concentrations (10, 7.5, 5, and 2.5 g/L). Biomass
suspensions were analyzed by Ribosomal Intergenic Spacer Analysis (RISA) to obtain
qualitative information on bacterial populations at each condition. Results showed that microbial
species composition responded more to influent glucose concentration than to HRT. Populations
detected at 2.5 g/L glucose concentration were taxonomically more diverse than at 5, 7.5, and 10
iv g/L glucose concentrations. The most intense band in RISA profiles at 10, 7.5, and 5 g/L glucose yielded sequences with high similarity (> 97%) to Clostridium acidisoli. At 2.5 g/L glucose, a Selenomonadaceae spp. was identified as being present in the reactor.
v Table of Contents Page
List of Tables………………………………………………………………………...... viii List of Figures……………………………………………………………………… …… ix Acknowledgments………………………………………………………………………. x
Chapter 1 Introduction……………………………………………………………………. 1 1.1 Problem Statements………………………………………………………………. 1 1.1.1 Perchlorate bioremediation………………………………………………… 1 1.1.2 Biological hydrogen generation ……………………………………………. 2 1.2 Organization of Dissertation……………………………………………………….. 3 1.3 References ……………………………………………………………………. 5
Chapter 2 Perchlorate reduction by a novel chemolithoautotrophic,hydrogen-oxidizing bacterium……………………………………………………………………………………10 2.1 Introduction………………………………………………………………………… 11 2.2 Results……………………………………………………………………………… 13 2.2.1 Enrichment and isolation…………………………………………………….. 13 2.2.2 Phenotypic characteristics…………………………………………………… 13 2.2.3 Autotrophic CO2 fixation by strain HZ……………………………………… 14 2.2.4 Phylogenetic analysis………………………………………………………… 14 2.3 Discussion………………………………………………………………………… 15 2.4 Experimental Procedures…………………………………………………………. 17 2.4.1 Medium and cultivation……………………………………………………… 17 2.4.2 Enrichment and isolation…………………………………………………….. 18 2.4.3 Electron microscopy……………………………………………..…………. 18 14 2.4.4 Determination of CO2 incorporation……………………………………… 19 2.4.5 Analytical techniques……………………………………………………….. 20 2.4.6 Phylogenetic analysis………………………………………………………… 20 2.5 References………………………………………………………………………… 22
Chapter 3 Molecular assessment of inoculated and indigenous bacteria in biofilms from a pilot- scale perchlorate-reducing bioreactor……………………………………………………… 30 3.1 Introduction………………………………………………………………………. 31 3.2 Materials and Methods……………………………………………………………. 34 3.3 Results…………………………………………………………………………….. 40 3.4 Discussion ……………………………………………………………………... 43 3.5 References………………………………………………………………………… 52
Chapter 4 Biological hydrogen production in a trickle-bed bioreactor……………………. 63 4.1 Introduction……………………………………………………………………….. 64 4.2 Materials and Methods…………………………………………………………… 65 4.2.1 Medium and culture conditions……………………………………… …….. 65 4.2.2 Reactor design and operation………………………………………… …… 66 4.2.3 Calculation…………………………………………………… …….……… 67 4.2.4 Determination of the reactor’s hydraulic retention time……....……………. 68
vi 4.2.5 Analytical procedures………………………………………………………. 68 4.3 Results and Discussion…………………………………………………………… 69 4.4 References………………………………………………………………………… 78
Chapter 5 Microbial community shifting as a function of hydraulic retention time (HRT) and substrate concentration in a chemostat H2-producing reactor……………………………... 81 5.1 Introduction……………………………………………………………………….. 81 5.2 Materials and Methods……………………………………………………………. 82 5.3 Results and Discussion…………………………………………………………… 84 5.4 References ………………………………………………………………………89
Appendix Data used to generate figures in chapter 2 to chapter 4………………………… 92
vii List of Tables
Table Page
Table 3.1 FISH probe sequences, target sites, formamide concentrations in the hybridization buffer and sodium chloride concentrations in the washing buffer……………………….. 51
Table 3.2 Phylogenetic summary of perchlorate-reducing community from cloning and sequencing results………………………………………………………………………… 52
Table 4.1 Summary of H2 production rates and conversion efficiency……………… 71
Table 4.2 Summary of volatile fatty acids production………………………………. 72
Table 4.3 Comparison of hydrogen production rates in the trickle-bed reactor with those reported using a CSTR……………………………………………………………………. 73
Table 5.1 Phylogenetic summary of hydrogen-producing community from cloning and sequencing results…………………………………………………………………………. 87
viii List of Figures
Figure Page
Figure 2.1 Perchlorate reduction by the chemolithoautotrophic hydrogen-oxidizing enrichment culture…………………………………………………………………………. 27
Figure 2.2 Scanning electron micrograph of anaerobically grown cells of strain HZ… 27
Figure 2.3 Growth of strain HZ with hydrogen as the electron donor and perchlorate (10 mM) as the electron acceptor with carbon dioxide as the only carbon source………… 28
Figure 2.4 Neighbor-joining phylogenetic tree of 16S rDNA sequences of strain HZ and others…………………………………………………………………………………... 29
Figure 3.1 Perchlorate concentration profile in the reactor with two flow rates……… 58
Figure 3.2 Chemical profiles in the reactor of acetate, oxygen, nitrate, and perchlorate at 0.34 L m-2s-1 flow rate……………………………………………………... 59
Figure 3.3 Inverted image of SYBR-green stained polyacrylamide gel showing ribosomal intergenic spacer profiles of DNA extracted from the reactor biofilms………... 60
Figure 3.4 FISH detection of Dechloromonas and Dechlorosoma species…………… 61
Figure 3.5 Relative abundance of bacteria hybridizing with EUB338, Monas, and Soma probes on the basis of total field area…………………………………………... 62
Figure 4.1 A picture and a schematic drawing of the trickle-bed reactor…………….. 74
Figure 4.2 Determination of reactor detention time by tracer study………………….. 75
Figure 4.3 Biogas production over time……………………………………………… 76
Figure 4.4 Headspace hydrogen concentrations over time…………………………… 76
Figure 4.5 Glucose, acetate, and butyrate profiles at the 3.3g/L glucose influent concentration……………………………………………………………………………… 77
Figure 5.1 Inverted image of ethidium bromide-stained agarose gels showing DNA bands in RISA profiles……………………………………………………………… 88
ix Acknowledgments
This publication was made possible in part by funding from National Science Foundation (BES9714575, BES0001900, BES0124674), the American Water Works Association Research Foundation (2557), and the Penn State Biogeochemical Research Initiative for Education (BRIE) sponsored by NSF (IGERT) grant DGE-9972759. I would like to thank my advisor Bruce Logan for giving me the freedom to work on interdisciplinary research topics, for encouraging me to ask important questions, and for numerous scientific discussions. He has been instrumental on every aspect of training me to become an independent researcher, from problem solving to scientific writing. I would like to thank my BRIE co-advisor Mary Ann Bruns for introducing me to molecular techniques, and for her constant encouragements throughout my dissertation research. I would like to thank Jay Regan for stimulating discussions and help on FISH experiments, Bill Burgos and Chris House for helpful suggestions. I would also like to thank Logan research group from January 2001 to August 2004, Bruns research group in the summers of 2001 to 2003, and Regan research group during the summer of 2003. Finally I would like to thank my wife Xin Luo for her love, support, after-school science discussions, and encouragement during my up and down times.
x Chapter 1
Introduction
1.1 Problem Statements
1.1.1 Perchlorate bioremediation
Perchlorate is a widespread contaminant found in groundwater and surface water environments (Logan 1998; Espenson 2000). Although perchlorate salts occur naturally only in some nitrate deposits in Chile (Schilt 1979; Espenson 2000), ammonium perchlorate has been manufactured in large quantities for use in explosives and solid rocket propellant. A new ion chromatography method with the minimum detection level to 2.5 to 4.0 ppb (depending on the water) led to the discovery of perchlorate in a large number of ground and surface water supplies
(Jackson et al. 2000). Recent studies have raised concerns that perchlorate, although used as a drug to treat thyrotoxicosis (Wolff 1998), may also inhibit the production of thyroid hormones in healthy populations. Based upon recent epidemiological and toxicological studies, the U. S.
Environmental Protection Agency's January 2002 draft report recommended a reference dose of
0.00003 mg kg-1 day-1. This dose would result in a drinking water standard of 1 µg L-1, using the
assumed factors of 70 kg body weight and 2 L of water consumption per day and a safety factor of 1000 (Renner 2003).
1 Due to perchlorate salts’ high solubility and nonreactivity in dilute solutions, biological treatment is considered to be a cost-effective means for large-scale perchlorate remediation
(Logan 2001). Perchlorate-reducing bacteria have been isolated (Rikken et al. 1996; Wallace et al. 1996; Bruce et al. 1999; Coates et al. 1999; Miller and Logan 2000; Giblin and Frankenberger
2001; Logan et al. 2001b; Waller et al. 2004), but all of them require organic carbon for energy and growth. In drinking water treatment systems, these organic compounds introduced into the water can lead to the release of unoxidized organic substrates that may stimulate regrowth in water distribution systems. There has been interest in bacteria that could be used in autotrophic reactors for drinking water treatment. Such systems have been used for nitrate removal (Gayle et al. 1989) but not yet for perchlorate degradation.
One issue that is important in engineered perchlorate treatment systems is whether the system should be bioaugmented. Engineered bioreactor systems have been designed and used to treat perchlorate-contaminated water (Wallace et al. 1998; Giblin et al. 2000; Miller and Logan
2000; Logan et al. 2001a; Logan and LaPoint 2002; Nerenberg and Rittmann 2002; Min et al.
2003), but there is little information about how well the bioaugmented microbes can compete with indigenous microbial members from the contaminated waters.
1.1.2 Biological hydrogen generation
Hydrogen gas holds the potential to be the next clean energy carrier. It produces water only when reacting with oxygen. Studies on biological hydrogen production has focused on using liquid-phase batch reactors (Okamoto et al. 2000; Van Ginkel et al. 2001; Logan et al.
2002; Nakashimada et al. 2002; Oh et al. 2003), or continuously stirred tank reactors (CSTR)
(Kataoka et al. 1997; Mizuno et al. 2000; Chen et al. 2001; Lay 2001; Hussy et al. 2003). In a
2 few cases, hydrogen generation from bacteria held on a packing in a column reactor has been examined, but only under saturated flow conditions (Rachman et al. 1998; Kumar and Das
2001). Conversion efficiencies reported in batch reactors and saturated packed bed reactors are typically less than 25%, based upon a maximum theoretical stoichiometry of 4 moles hydrogen from 1 mole of hexose. While stirring can facilitate hydrogen gas release from the liquid phase in a CSTR, it also consumes considerable electric power. To our knowledge there have been no reports on using trickle-bed reactors for biological hydrogen production. Trickle-bed reactors have been widely used in aerobic wastewater treatment. They do not require mechanical mixing power to operate and are relatively easy to construct.
Pure cultures of bacteria have been used to produce H2 (Taguchi et al. 1996; Kataoka et
al. 1997; Rachman et al. 1997; Chin et al. 2003). For practical applications using organic matter
from wastewaters, however, aseptic conditions are not possible so that mixed consortia must be
used. Selecting and maintaining a microbial consortium is key for stable operation of a H2- producing bioreactor. There are a few reports on microbial communities that developed in H2-
producing reactors. In all these studies, sequences closely related to Clostridium spp. have been
reported (Ueno et al. 2001; Fang et al. 2002; Oh et al. 2004). Except for one case (Oh et al.
2004), clone library or denaturing gradient gel electrophoresis (DGGE) was used to identify
community compositions.
1.2 Organization of Dissertation
During my dissertation, I worked on the two topics of perchlorate bioreduction and biohydrogen generation. Therefore this dissertation is organized into separate chapters based on manuscripts that I have written during the course of my studies. Chapter 2 is a manuscript on the
3 isolation of an autotrophic perchlorate-reducing bacterium. The hypothesis was that a pure culture can be obtained using the dilution-to-extinction method. I isolated Dechloromonas sp. strain HZ. Strain HZ is a facultative anaerobe that was demonstrated to be capable of growth solely on CO2 with hydrogen as electron donor and perchlorate as electron acceptor. Strain HZ
can also use oxygen, chlorate, and nitrate as electron acceptors under autotrophic conditions.
This manuscript has been published in Environmental Microbiology, entitled “Perchlorate
reduction by a novel chemolithoautotrophic, hydrogen-oxidizing bacterium” by Zhang H, Bruns
MA, and Logan BE. I designed the experiments based on discussions with Dr. Logan on
dilution techniques, and Dr. Bruns on molecular identification. I performed the experiments and wrote the manuscript which was edited by Dr. Logan and Dr. Bruns.
Chapter 3 was a manuscript on analyzing the microbial communities in a bioreactor using
molecular techniques. It was not certain whether Dechlorosoma sp. strain KJ, the inoculum used
in a pilot-scale, packed-bed reactor for perchlorate treatment, will persist after the reactor was
operated for 6 months. Polymerase chain reaction (PCR)–based fingerprinting experiments
failed to detect the inoculum from total DNA extractions. Fluorescent In Situ Hybridization
(FISH) gave a more complete picture: low numbers of Dechlorosoma spp. were in the inner
biofilms, while Dechloromonas spp. were in the shallow portions of the biofilms. This
manuscript, entitled “Molecular assessment of inoculated and indigenous bacteria in biofilms
from a pilot-scale perchlorate-reducing bioreactor” by Zhang H, Logan BE, Regan JM,
Achenbach LA, and Bruns MA, has been accepted and is in press in Microbial Ecology. I
designed the experiments for this study based on discussions with Dr. Bruns, Dr. Logan, and Dr.
Regan. I performed all the experiments and wrote the first draft of the manuscript. Dr. Bruns
and Dr. Logan edited the manuscript.
4 Chapter 4 represents a manuscript in preparation for submission to Applied Microbiology and Biotechnology, entitled “Biological hydrogen production in a trickle-bed reactor” by Zhang
H, Bruns MA, and Logan BE. The hypothesis was that high concentrations of hydrogen gas can be generated in a trickle-bed reactor with no mixing needed. I found that hydrogen was produced at a rate higher or comparable to those reported using CSTR reactors. The gas-phase hydrogen concentrations obtained in this study were 74-79%. The experiments were designed by me based on discussions with Dr. Logan. I performed the experiments and wrote the manuscript with suggestions and manuscript editing by Dr. Logan and Dr. Bruns.
Chapter 5 describes my contribution to a collaborative investigation of microbial community composition in a fermentative H2-producing reactor as a function of two operational
parameters: hydraulic retention time (HRT) and substrate loading rate. The hypothesis was that
spore-forming bacteria such as Clostridium spp., will be in the reactor. Another hypothesis was that the composition of microbial species would be affected by HRT and substrate loading
reflecting different growth rates and nutrient requirements. The results showed microbial species
that are closely related to Clostridium acidisoli were present in all combinations of HRT and
organic loading except for the lowest organic loading used in this study. I performed the
experiments, analyzed the results, and wrote the chapter with suggestions from Dr. Logan and
Dr. Bruns. This chapter is not yet submitted. The results will be incorporated into a future
manuscript.
1.3 References
Bruce RA, Achenbach LA, Coates JD (1999) Reduction of (per)chlorate by a novel organism
isolated from paper mill waste. Environ Microbiol 1:319-329
5 Chen CC, Lin CY, Chang JS (2001) Kinetics of hydrogen production with continuous anaerobic
cultures utilizing sucrose as the limiting substrate. Appl Microbiol Biotechnol 57:56-64
Chin HL, Chen ZS, Chou CP (2003) Fedbatch operation using Clostridium acetobutylicum
suspension culture as biocatalyst for enhancing hydrogen production. Biotechnol Prog
19:383-388
Coates JD, Michaelidou U, Bruce RA, O'Connor SM, Crespi JN, Achenbach LA (1999)
Ubiquity and diversity of dissimilatory (per)chlorate-reducing bacteria. Applied and
Environmental Microbiology 65:5234-5241
Espenson JH (2000) The problem and perversity of perchlorate. In: Urbansky ET (ed)
Perchlorate in the environment. Kluwer Academic/Plenum, New York, pp 1-8
Fang HH, Zhang T, Liu H (2002) Microbial diversity of a mesophilic hydrogen-producing
sludge. Appl Microbiol Biotechnol 58:112-118
Gayle BP, Boardman GD, Sherrard JH, Benoit RE (1989) Biological denitrification of water. J
Environ Eng-ASCE 115:930-943
Giblin T, Frankenberger WT (2001) Perchlorate and nitrate reductase activity in the perchlorate-
respiring bacterium perclace. Microbiol Res 156:311-315
Giblin T, Herman D, Deshusses MA, Frankenberger WT (2000) Removal of perchlorate in
ground water with a flow-through bioreactor. J Environ Qual 29:578-583
Hussy I, Hawkes FR, Dinsdale R, Hawkes DL (2003) Continuous fermentative hydrogen
production from a wheat starch co-product by mixed microflora. Biotechnol Bioeng
84:619-626
6 Jackson PE, Gokhale S, Rohrer JS (2000) Recent developments in the analysis of perchlorate
using ion chromatography. In: Urbansky ET (ed) Perchlorate in the environment. Kluwer
Academic/Plenum, New York, pp 37-44
Kataoka N, Miya A, Kiriyama K (1997) Studies on hydrogen production by continuous culture
system of hydrogen-producing anaerobic bacteria. Water Sci Technol 36:41-47
Kumar N, Das D (2001) Continuous hydrogen production by immobilized Enterobacter cloacae
iit-bt 08 using lignocellulosic materials as solid matrices. Enzyme Microb Tech 29:280-
287
Lay JJ (2001) Biohydrogen generation by mesophilic anaerobic fermentation of microcrystalline
cellulose. Biotechnol Bioeng 74:280-287
Logan BE (1998) A review of chlorate- and perchlorate-respiring microorganisms. Bioremed J
2:69-79
Logan BE (2001) Assessing the outlook for perchlorate remediation. Environ Sci Technol
35:482a-487a
Logan BE, Kim K, Price S (2001a) Perchlorate degradation in bench- and pilot scale ex-situ
bioreactors. In: Leeson A, Peyton BM, Means JL, Magar VS (eds) Bioremediation of
inorganic compounds. Battelle Press, Columbus, OH, pp 303-308
Logan BE, LaPoint D (2002) Treatment of perchlorate- and nitrate-contaminated groundwater in
an autotrophic, gas phase, packed-bed bioreactor. Water Res 36:3647-3653
Logan BE, Oh S, Kim IS, Van Ginkel S (2002) Biological hydrogen production measured in
batch anaerobic respirometers. Environ Sci Technol 36:2530-2535
Logan BE, Zhang H, Mulvaney P, Milner MG, Head IM, Unz RF (2001b) Kinetics of
perchlorate- and chlorate-respiring bacteria. Appl Environ Microbiol 67:2499-2506
7 Miller JP, Logan BE (2000) Sustained perchlorate degradation in an autotrophic, gas-phase,
packed-bed bioreactor. Environ Sci Technol 34:3018-3022
Min B, Evans PJ, Chu A, Logan BE (2003) Perchlorate removal in sand and plastic media
bioreactors. Water Res In press
Mizuno O, Dinsdale R, Hawkes FR, Hawkes DL, Noike T (2000) Enhancement of hydrogen
production from glucose by nitrogen gas sparging. Bioresource Technol 73:59-65
Nakashimada Y, Rachman MA, Kakizono T, Nishio N (2002) Hydrogen production of
Enterobacter aerogenes altered by extracellular and intracellular redox states. Int J
Hydrogen Energ 27:1399-1405
Nerenberg R, Rittmann BE (2002) Perchlorate as a secondary substrate in a denitrifying hollow-
fiber membrane biofilm reactor. Water Sci Technol 2:259?65
Oh SE, Iyer P, Bruns MA, Logan BE (2004) Biological hydrogen production using a membrane
bioreactor. Biotechnol Bioeng 87:119-127
Oh SE, Van Ginkel S, Logan BE (2003) The relative effectiveness of ph control and heat
treatment for enhancing biohydrogen gas production. Environ Sci Technol 37:5186-5190
Okamoto M, Miyahara T, Mizuno O, Noike T (2000) Biological hydrogen potential of materials
characteristic of the organic fraction of municipal solid wastes. Water Sci Technol 41:25-
32
Rachman MA, Furutani Y, Nakashimada Y, Kakizono T, Nishio N (1997) Enhanced hydrogen
production in altered mixed acid fermentation of glucose by Enterobacter aerogenes. J
Ferment Bioeng 83:358-363
8 Rachman MA, Nakashimada Y, Kakizono T, Nishio N (1998) Hydrogen production with high
yield and high evolution rate by self-flocculated cells of Enterobacter aerogenes in a
packed-bed reactor. Appl Microbiol Biot 49:450-454
Renner R (2003) Environmental health. Academy to mediate debate over rocket-fuel
contaminants. Science 299:1829
Rikken GB, Kroon AGM, vanGinkel CG (1996) Transformation of (per)chlorate into chloride by
a newly isolated bacterium: Reduction and dismutation. Appl Microbiol Biot 45:420-426
Schilt AA (1979) Perchloric acid and perchlorates. G. F. Smith Chemical Co., Columbus, Ohio
Taguchi F, Yamada K, Hasegawa K, TakiSaito T, Hara K (1996) Continuous hydrogen
production by Clostridium sp strain no. 2 from cellulose hydrolysate in an aqueous two-
phase system. J Ferment Bioeng 82:80-83
Ueno Y, Haruta S, Ishii M, Igarashi Y (2001) Microbial community in anaerobic hydrogen-
producing microflora enriched from sludge compost. Appl Microbiol Biotechnol 57:555-
562
Van Ginkel S, Sung SW, Lay JJ (2001) Biohydrogen production as a function of pH and
substrate concentration. Environ Sci Technol 35:4726-4730
Wallace W, Beshear S, Williams D, Hospadar S, Owens M (1998) Perchlorate reduction by a
mixed culture in an up-flow anaerobic fixed bed reactor. J Ind Microbiol Biot 20:126-131
Wallace W, Ward T, Breen A, Attaway H (1996) Identification of an anaerobic bacterium which
reduces perchlorate and chlorate as Wolinella succinogenes. J Ind Microbiol 16:68-72
Waller AS, Cox EE, Edwards EA (2004) Perchlorate-reducing microorganisms isolated from
contaminated sites. Environ Microbiol 6:517-527
Wolff J (1998) Perchlorate and the thyroid gland. Pharmacol Rev 50:89-106
9 Chapter 2
Perchlorate reduction by a novel chemolithoautotrophic,hydrogen-oxidizing bacterium
Abstract
Water treatment technologies are needed that can remove perchlorate from drinking water without introducing organic chemicals that can stimulate bacterial growth in water distribution systems. Hydrogen is an ideal energy source for bacterial degradation of perchlorate as it leaves no organic residue and is sparingly soluble. We describe here the isolation of a perchlorate- respiring, hydrogen-oxidizing bacterium (Dechloromonas sp. strain HZ) that grows with carbon dioxide as sole carbon source. Strain HZ is a gram-negative, rod-shaped facultative anaerobe that was isolated from a gas-phase autotrophic packed-bed biofilm reactor treating perchlorate- contaminated groundwater. The ability of strain HZ to grow autotrophically with carbon dioxide as the sole carbon source was confirmed by demonstrating that biomass carbon (100.9%) was
14 derived from CO2. Chemolithotrophic growth with hydrogen was coupled with complete
reduction of perchlorate (10 mM) to chloride with a maximum doubling time of 8.9 hours.
Strain HZ also grew using acetate as the electron donor and chlorate, nitrate, or oxygen (but not sulfate) as an electron acceptor. Phylogenetic analysis of the 16S rRNA sequence placed strain
HZ in the genus Dechloromonas within the β subgroup of the Proteobacteria. The study of this
and other novel perchlorate-reducing bacteria may lead to new, safe technologies for removing
perchlorate and other chemical pollutants from drinking water.
10 2.1 Introduction
Perchlorate salts are manufactured in large quantities for use as oxidizers in solid rocket propellants, explosives, and automobile air bag inflators (Espenson, 2000; Logan, 2001).
Although it reacts energetically when dry, perchlorate in aqueous solution is extremely stable and highly soluble. Until 1997, perchlorate could not be measured in water below 100 ppb (1
µM). A new ion chromatography procedure reduced the minimum detection level to 2.5 to 4.0 ppb (depending on the water) and led to the discovery of perchlorate in a large number of ground and surface water supplies (Jackson et al., 2000; Logan, 2001). Recent epidemiological studies have recommended a lower perchlorate maximum contaminant level that, if adopted by the USEPA using standard safety factors, could result in a drinking water standard of 1 ppb
(Anonymous, 2002). This is a concentration measurable only with more sensitive and expensive techniques than ion chromatography (Handy et al., 2000; Koester et al., 2000). The California
Department of Health Services has reduced their Action Level for perchlorate from 18 ppb to 4 ppb in response to the new health study.
Although perchlorate is known to occur naturally only in some nitrate deposits in Chile
(Schilt, 1979; Espenson, 2000), bacteria capable of degrading perchlorate are surprisingly widespread in nature (Logan, 1998; Coates et al., 1999; Logan et al., 2001b; Wu et al., 2001).
Despite their abundance, only a few perchlorate-reducing bacteria (PRB) have been physiologically and phylogenetically characterized (Wallace et al., 1996; Bruce et al., 1999;
Achenbach et al., 2001; Logan et al., 2001b). Kinetic constants (µm and KS) have been
determined for two isolates (Dechlorosoma sp. KJ and Dechlorosoma sp. PDX) under different
electron-accepting conditions (Logan et al., 2001b), and pilot-scale bioreactors have been
11 designed and inoculated with a pure culture of Dechlorosoma sp. KJ for treatment of perchlorate and nitrate contaminated groundwater (Logan et al., 2001a; Evans et al., 2002).
So far, all perchlorate-degrading isolates reported to date have been heterotrophic
(Logan, 1998; Achenbach et al., 2001), and therefore require organic substrates for synthesizing cellular materials. Growth substrates introduced into the water to support the growth of heterotrophic PRB In drinking water treatment systems can lead to the release of unoxidized organic substrates that may stimulate subsequent microbiological growth in water distribution systems. Bioreactors that use only hydrogen gas, carbon dioxide, and inorganic nutrients, are being investigated for treatment of perchlorate-contaminated groundwater (Giblin et al., 2000;
Miller and Logan, 2000; Logan and LaPoint, 2002; Nerenberg and Rittmann, 2002). However, perchlorate-reducing isolates that grow solely on inorganic energy and carbon sources have not been previously isolated from these reactors. In one case an isolate from an autotrophic reactor
(Dechloromonas sp. strain JM) was found to reduce perchlorate with hydrogen, but it needed an organic substrate such as acetate for growth, and could not fix CO2 (Miller and Logan, 2000). In
another case a consortium of four different bacteria was found to be necessary for perchlorate degradation with hydrogen (Giblin et al., 2000). These findings have led to speculation that several bacterial species acting in concert are required to degrade perchlorate in autotrophic bioreactors (Giblin et al., 2000).
Here we report on the isolation of a novel chemolithoautotrophic, perchlorate-reducing, hydrogen-oxidizing bacterium (Dechloromonas sp. strain HZ) using a dilution-to-extinction technique. Autotrophic growth on CO2 was confirmed by demonstration of quantitative
14 incorporation of CO2 into cell mass.
12 2.2 Results
2.2.1 Enrichment and isolation
After 28 days of incubation with H2 and CO2, perchlorate reduction (3 mM) was
observed by the enrichment culture from the bioreactor. Successive transfers (three times, 5%
inoculum) to fresh H2-CO2-perchlorate medium reduced the time for complete (< 0.01 mM)
perchlorate removal to only two days (Fig. 1.1). Serial dilution to extinction produced a culture
with morphologically uniform cells. While this culture could be repeatedly transferred in liquid
medium, there was no growth on agar (Difco) plates containing the same medium and a gas
mixture of H2 and CO2. Because it was not possible to grow and isolate colonies on plates, the
purity of the culture was initially evaluated only by microscopic examination. Molecular tests to
confirm purity were based on extracting genomic DNA from the culture for PCR amplification
of the variable 16S-23S ribosomal RNA spacer region and determining that independently
cloned PCR products had identical DNA sequences.
2.2.2 Phenotypic characteristics
Cells of strain HZ are Gram-negative rods 0.3 × 1.8 µm (Fig. 1.2) capable of complete
reduction of perchlorate to chloride under H2-oxidizing, autotrophic growth conditions (Fig. 1.3).
The observed maximum doubling time was 8.9 h. No growth was detected when hydrogen,
perchlorate, or CO2 was omitted from the medium. Methane was not detected during autotrophic
growth. Nitrate and chlorate, but not sulfate, could also serve as electron acceptors under
autotrophic growth conditions. Heterotrophic growth was successful with acetate (20 mM) as the
13 electron donor and perchlorate (10 mM), chlorate (10 mM), nitrate (10 mM), or oxygen (21 kPa) as electron acceptors. The ability to use nitrate and chlorate as the electron acceptors is a common characteristic of many heterotrophic perchlorate-reducing organisms (Logan, 1998;
Achenbach et al., 2001).
2.2.3 Autotrophic CO2 fixation by strain HZ
14 The autotrophic growth of strain HZ was confirmed by measuring incorporation of CO2 into cellular material (all measurements in triplicate). After 48 h incubation, the cell dry weight was 147.9 ± 12.3 µg/ml. The carbon content of nonlabeled cells grown under the same conditions (independently determined to be 47 ± 1% carbon) was used to calculate a biomass carbon dry weight of 69.7 ± 7.3 µg-C/ml. Mean weight of incorporated C (70.3 ± 15.3 µg-C/ml) was calculated by dividing the total activity of cells recovered from 1 ml of culture by the
14 specific activity of CO2 per µg-C in the gas phase (see Experimental procedures). Thus, a total
of 100.9% (70.3/69.7×100%) of carbon in the cell biomass could be attributed to fixation of CO2.
Cells cultured without hydrogen (under N2/CO2) or killed with formaldehyde (2%) did not show
14 significant CO2 uptake (0.014% and 0.012% uptake of total radiolabel, respectively, as opposed
to 0.93% by cells grown under H2/CO2).
2.2.4 Phylogenetic analysis
Phylogenetic analysis based on the nearly full-length 16S rRNA sequence was used to classify strain HZ as a Dechloromonas sp. A phylogenetic tree (Fig. 1.4) inferred from distance
analysis of 16S rRNA sequence data reveals that strain HZ is one of a several new isolates in the
14 β subclass of Proteobacteria that can completely reduce perchlorate to chloride. Strain HZ falls within the Dechloromonas “clade” of the Rhodocyclus group but is distinguished physiologically from other isolates by its ability to grow autotrophically with hydrogen as the electron donor.
Although others have identified perchlorate degraders within the α, β, and ε subclasses of
Proteobacteria, most isolates (ca. 70%) fall within the Dechloromonas and Dechlorosoma clades of the β subclass, and are monophyletic (Coates et al., 1999). Both clades, however, contain representatives that do not degrade perchlorate, such as Ferribacterium limneticum and
Rhodocyclus spp. Bootstrap values for Fig. 1.4 support a more distant relationship between
Dechloromonas and Dechlorosoma spp. than between Dechlorosoma, Rhodocyclus,
Propionibacter, and Propionivibrio spp., indicating that perchlorate degradation ability is not a defining physiological characteristic of bacteria in these two clades.
2.3 Discussion
The closest relative to strain HZ is Dechloromonas sp. strain JM, a heterotrophic bacterium having 99.8% similarity in its full-length 16S rRNA. The 16S-23S ribosomal intergenic spacers, that are highly variable among bacteria, have identical lengths in strains HZ and JM (447 base pairs), as well as 97.6% sequence similarity. Despite a close phylogenetic relationship between strains HZ and JM, it is known that bacteria with >99% similar 16S rRNA gene sequences can exhibit 30-70% dissimilarity across their complete genomes (Stackebrandt and Gobel, 1994). The dissimilar portions of strain HZ’s genome could therefore contain the genes for its autotrophy-associated enzymes, or such genes could reside on extrachromosomal megaplasmids (Stouthamer and Kooijman, 1993). Autotrophic and nonautotrophic
15 representatives have also been observed among subspecies of Paracoccus denitrificans (Jordan et al., 1997).
In addition to respiration using perchlorate, a chemical only introduced into the environment relatively recently, Dechloromonas sp. strain HZ and other perchlorate- and chlorate-respiring bacteria harbor other unique physiological properties such as their hydrogen oxidizing capabilities and novel respiratory enzymes. These bacteria all contain chlorite dismutase, an enzyme capable of disproportionating chlorite to chloride and oxygen with high efficiency (van Ginkel et al., 1996). This enzyme can be used in reactors for the removal of chlorate produced from drinking water disinfection with chlorine dioxide (van Ginkel et al.,
1998). The first microbe capable of growth on benzene under denitrifying conditions was a perchlorate-respiring bacterium (Coates et al., 2001).
Hydrogen gas reactors have been used in the past for nitrate removal via denitrification from drinking water (Gayle et al., 1989). New reactor designs using packed beds and hollow fiber membranes are also being researched for treating perchlorate-contaminated water. The main advantage to using hydrogen is that it is only sparingly soluble in water (1.62 mg/L at 25
o C and 1 atm H2) (Chapelle et al., 1997). The use of this inorganic energy source in a bioreactor
reduces the potential for contamination of water distribution systems by growth of bacteria on
non-oxidized substrate. Hydrogen gas-based drinking water treatment systems may offer other
advantages for treating drinking water. At many sites, chlorinated aliphatic compounds, such as
trichloroethylene (TCE), are common contaminants of perchlorate-contaminated groundwater
(Catts, 1999). Although chemicals such as TCE may persist in the subsurface environment,
these chemicals can be degraded by halorespiring bacteria using hydrogen (Fennell et al., 1997;
Maymo-Gatell et al., 1997). Thus, it may be possible to use mixtures of specific bacteria such as
16 Dechloromonas sp. strain HZ in hydrogen-gas reactors to treat water contaminated with perchlorate and other common groundwater pollutants.
2.4 Experimental procedures
2.4.1 Medium and cultivation
The enrichment medium (Miller and Logan, 2000) was prepared using ultrapure water
(Milli-Q system; MilliPore Corp.) and contained (per liter): K2HPO4, 3 g; NaH2PO4•H2O, 1.7g;
NH4H2PO4, 1 g; NaClO4, 0.62 g; NaHCO3, 1 g; MgSO4•7H2O, 50 mg; Na2EDTA, 6 mg;
CaCl2•2H2O, 1 mg; Na2MoO4•2H2O, 0.2 mg; CoCl2•6H2O, 0.4 mg; Na2SeO3•5H2O, 0.1 mg;
NiCl2•6H2O, 0.1; ZnSO4•7H2O, 2; FeSO4•7H2O, 5 mg; CuCl2•2H2O 0.2 mg; MnCl2•4H2O, 1 mg; and H3BO3, 0.6 mg. After being degassed overnight in an anaerobic chamber (Coy Scientific
Products), serum bottles (100 ml) containing the anaerobic medium were closed with thick butyl
rubber stoppers, sealed with aluminum caps, and removed from the glove box. The headspace of
each bottle was exchanged with a mixture of H2:CO2 (80:20, v:v) three times to 200 kPa
overpressure. The medium was sterilized by autoclaving at 121 oC for 30 min. The final pH of
the cooled media was 6.8. All transfers were made in the anaerobic chamber. Cultures were
incubated in the dark at 28 oC on a rotary shaker at 200 rpm.
17 2.4.2 Enrichment and isolation
A cell suspension (2.5 ml) from a perchlorate-degrading, autotrophic laboratory packed- bed reactor (Miller and Logan, 2000) was added to the organic-free medium and incubated in the dark at 28 °C. After perchlorate reduction (3 mM) was observed, repeated subcultivation (5% transfers) was carried out to further enrich the chemolithoautotrophic perchlorate-reducing bacteria. A serial dilution of the enrichment culture to 10-3 produced growth after eight weeks.
Following several successive transfers from this culture, a second serial dilution was performed.
A pure culture was obtained from the lowest positive (10-5) dilution. Purity was initially
examined using a microscope, but it was confirmed by examining the intergenic spacer
sequences from multiple clones (see below).
2.4.3 Electron microscopy
Anaerobically grown cells were filtered onto 0.22 µm polycarbonate filters and fixed
using 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer for one hour. Filters were rinsed in
the same buffer, fixed with 1% osmium tetroxide (in 0.1 M sodium cacodylate buffer),
dehydrated using a concentration gradient series of ethanol, critical point dried in a BAL-TEC
SCD 030 CPD, sputter-coated with gold/palladium in a BAL-TEC 050 coater (Techno Trade,
Manchester, NH), and viewed with a JEOL JSM 5400 scanning electron microscope at 20 kV
(JEOL, Peabody, MA).
18 14 2.4.4 Determination of CO2 incorporation
Bacterial fixation of carbon dioxide was calculated using a medium containing
radiolabeled carbon dioxide by: (i) calculating the fraction of carbon dioxide in the container that
was radiolabeled (via the specific activity), and (ii) measuring the amount of radiolabel
incorporated into the bacteria. Anaerobic medium (20 ml) containing NaHCO3 was prepared as
14 described above, and amended with NaH CO3 (0.5 ml, 336 kBq/ml). To ensure equilibration of
the radiolabel between the gas and liquid phases, the bottle was shaken at 200 rpm for 8 hours.
Because bottles contained radiolabeled and non-radiolabeled CO2 in an unknown proportion, gas
14 samples were analyzed to determine the specific activity of CO2 in the bottle using the method
of Brysch et al. (1987). Briefly, gas (200 µl) was withdrawn using a gastight syringe
(SamplelockTM; Hamilton Company) and injected into a sealed vial containing NaOH (2 ml, 1
M) to trap the CO2. A portion of the liquid in the CO2 trap (0.5 ml) was mixed with 10 ml
scintillation cocktail (Hionic-FluorTM, Packard Instrument Company) and the 14C concentration
(L as Bq/ml) determined by liquid scintillation counting (Rackbeta 1217, LKB Wallac). The total
concentration (K as µg/ml) of carbon in the CO2 trap was determined gravimetrically by BaCO3 precipitation using a 0.2 µm membrane filter and a microbalance (Mettler Toledo UMT2).
Because the mass of radiolabel and mass of total carbon dioxide are now known, the specific
14 activity, S, of CO2, could be calculated as S = L/K = 1.12 ± 0.10 Bq/µg-C (mean ± SD).
To determine biomass carbon derived from CO2, the radiolabel incorporated into the cells
was used to calculate the carbon fixation from the specific activity. Cells (2.5%) were transferred
14 into the medium containing NaH CO3 and incubated for 48 h. Killed controls (2%
formaldehyde) were used to determine adsorption of the radiolabel. Samples (1 ml) were
14 removed and mixed with 6 M phosphoric acid (0.1 ml) to allow the escape of unfixed CO2
19 (Jordan et al., 1997), filtered through 0.22 µm membrane filters (Osmonics Corp.), washed three times with distilled water (5 ml), and added to the scintillant for determination of the 14C content.
Biomass carbon derived from CO2 (designated as BC) was calculated as 70.3 ± 15.3 µg-C/ml
using the equation BC = A/S, where A is the total activity of cells recovered from 1 ml culture
(78.5 ± 10.3 Bq/ml), and S the specific activity of CO2-C in the headspace as determined above
(1.12 ± 0.10 Bq/µg-C). To examine the radiolabel recovery during the experiments, both gas
phase and aqueous phase samples (200 µl) were taken for determination of total 14C radioactivity
before and after the experiments.
2.4.5 Analytical techniques
Chloride and perchlorate were analyzed using ion chromatography, and cell dry weight
(DW) was measured using a microbalance as previously described (Logan et al., 2001b). Cell carbon content was determined as a percent of dry weight by using a Fisons NA1500 Elemental
Analyzer equipped with an IRMS detector (Pella, 1990).
2.4.6 Phylogenetic analysis
Genomic DNA was extracted (Qiagen) from cell pellets of strain HZ for polymerase chain reactions using the rDNA primers (Lane, 1991) to obtain 16S rRNA and 16S-23S spacer sequences (Bruns et al., 2001). PCR products were cloned using the TOPO TA cloning kit
(Invitrogen), sequenced (ABI), and edited with EditSeq and SeqMan programs (Lasergene). The nearly full-length 16S rRNA sequence for strain HZ was manually aligned using secondary
20 structure information with related sequences downloaded from GenBank and the rRNA WWW
Server (HTTP://RRNA.UIA.AC.BE/). Aligned sequence data, corresponding to E. coli rRNA
positions 190-1392 (Thompson et al., 1997), were subjected to bootstrap resampling (100
replicates) using SEQBOOT from the PHYLIP package (Felsenstein, 1993). Distance,
maximum-likelihood and parsimony analyses based on random-sequence additions were carried
out with the DNADIST, DNAML, and DNAPARS programs, respectively, of PHYLIP, and
consensus trees were generated with the CONSENSE program. Paracoccus denitrificans, a
representative of α-Proteobacteria, was used as outgroup. Consistent topologies and similar
bootstrap values were obtained for all three types of trees. GenBank accession numbers for
sequences used in this analysis are: Paracoccus denitrificans IAM12479, Y17512; Azoarcus
evansii, X77679; A. tolulyticus strain 4FB10, AF229876; Thauera selenatis, Y17591; T.
terpenica strain 21Mol, AJ005818; Azovibrio sp. strain BS20-3, AF011349; Az. restrictus strain
BS1-14, AF011348; Rhodocyclus tenuis, D16209; R. purpureus, M34132; Propionivibrio
dicarboxylicus, Y17601; Propionibacter pelophilus, AF016690; Dechlorosoma sp. strain KJ,
AF323491; Dechlorosoma suillum, AF170348; Dechlorosoma sp. strain PDX, AF323490;
AF016690; Ferribacterium limneticum, Y17060; Dechloromonas sp. strain MissR, AF170357;
Dechloromonas sp. strain SIUL, AF170356; Dechloromonas sp. strain RCB, AY032610;
Dechloromonas sp. strain JJ, AY032611; Dechloromonas sp. strain FL2, AF288771;
Dechloromonas sp. strain FL9, AF288773; Dechloromonas agitata, AF047462; Dechloromonas
sp. isolate CL, AF170354; Dechloromonas sp. strain JM, AF323489; perchlorate-reducing
bacterium MLC33, AF444791; Dechloromonas sp. strain HZ, AF479766.
21 Acknowledgments
We thank Chris House for advice and providing gassing equipment, Jianlin Xu for help with heterotrophic growth tests, Denyce Matlin for conducting cell carbon analysis, Rosemary Walsh of the Electron Microscopy Facility in the Life Science Consortium at Penn State University, and
Dan Hannon for technical assistance. This research was funded by the National Science
Foundation (Grants DGE-9972759, BES9714575, BES0001900) and the American Water Works
Association Research Foundation (2557). H.Z. was supported by the Penn State Biogeochemical
Research Initiative for Education (BRIE) sponsored by NSF (IGERT) grant DGE-9972759. The
GenBank accession number for Dechloromonas sp. strain HZ is AF479766.
2.5 References
Achenbach, L., Michaelidou, U., Bruce, R., Fryman, J., and Coates, J. (2001) Dechloromonas
agitata gen. nov., sp. nov. and Dechlorosoma suillum gen. nov., sp. nov., two novel
environmentally dominant (per)chlorate-reducing bacteria and their phylogenetic position.
Int J Syst Evol Microbiol 51: 527-533.
Anonymous (2002) Low-level perchlorate exposures. Environ Sci Technol 36: 125 A.
Bruce, R.A., Achenbach, L.A., and Coates, J.D. (1999) Reduction of (per)chlorate by a novel
organism isolated from paper mill waste. Environ Microbiol 1: 319-329.
Bruns, M.A., Hanson, J.R., Mefford, J., and Scow, K.M. (2001) Isolate PM1 populations are
dominant and novel methyl tert-butyl ether-degrading bacterial in compost biofilter
enrichments. Environ Microbiol 3: 220-225.
22 Brysch, K., Schneider, C., Fuchs, G., and Widdel, F. (1987) Lithoautotrophic growth of sulfate-
reducing bacteria, and description of Desulfobacterium autotrophicum gen-nov, sp-nov. Arch
Microbiol 148: 264-274.
Catts, J.G. (1999) Biochemical removal of perchlorate from San Gabriel groundwater and
potable use of the treated water. Abstr Pap - Am Chem Soc 218: pp. 107-109.
Chapelle, F.H., Vroblesky, D.A., Woodward, J.C., and Lovley, D.R. (1997) Practical
considerations for measuring hydrogen concentrations in groundwater. Environ Sci Technol
31: 2873-2877.
Coates, J.D., Michaelidou, U., Bruce, R.A., O'Connor, S.M., Crespi, J.N., and Achenbach, L.A.
(1999) Ubiquity and diversity of dissimilatory (per)chlorate-reducing bacteria. Appl Environ
Microbiol 65: 5234-5241.
Coates, J.D., Chakraborty, R., Lack, J.G., O'Connor, S.M., Cole, K.A., Bender, K.S., and
Achenbach, L.A. (2001) Anaerobic benzene oxidation coupled to nitrate reduction in pure
culture by two strains of Dechloromonas. Nature 411: 1039-1043.
Espenson, J.H. (2000) The problem and perversity of perchlorate. In Perchlorate in the
environment. Urbansky, E.T. (ed). New York: Kluwer Academic/Plenum, pp. 1-8.
Evans, P., Chu, A., Liao, S., Price, S., Min, B., and Logan, B.E. (2002) Pilot testing of a
bioreactor for perchlorate-contaminated groundwater treatment. In Third International
Conference on Remediation of Chlorinated and Recalcitrant Compounds. Monterey, CA In
press.
Felsenstein, J. (1993) PHYLIP - Phylogeny Inference Package. University of Washington,
Seattle, WA.
23 Fennell, D.E., Gossett, J.M., and Zinder, S.H. (1997) Comparison of butyric acid, ethanol, lactic
acid, and propionic acid as hydrogen donors for the reductive dechlorination of
tetrachloroethene. Environ Sci Technol 31: 918-926.
Gayle, B.P., Boardman, G.D., Sherrard, J.H., and Benoit, R.E. (1989) Biological denitrification
of water. J Environ Eng-ASCE 115: 930-943.
Giblin, T.L., Herman, D.C., and Frankenberger, W.T. (2000) Removal of perchlorate from
ground water by hydrogen-utilizing bacteria. J Environ Qual 29: 1057-1062.
Handy, R., Barnett, D.A., Purves, R.W., Horlick, G., and Guevremont, R. (2000) Determination
of nanomolar levels of perchlorate in water by ESI-FAIMS-MS. J anal at spectrom 15: 907-
911.
Jackson, P.E., Gokhale, S., and Rohrer, J.S. (2000) Recent developments in the analysis of
perchlorate using ion chromatography. In Perchlorate in the environment. Urbansky, E.T.
(ed). New York: Kluwer Academic/Plenum, pp. 37-44.
Jordan, S.L., McDonald, I.R., Kraczkiewicz-Dowjat, A.J., Kelly, D.P., Rainey, F.A., Murrell,
J.C., and Wood, A.P. (1997) Autotrophic growth on carbon disulfide is a property of novel
strains of Paracoccus denitrificans. Arch Microbiol 168: 225-236.
Koester, C.J., Beller, H.R., and Halden, R.U. (2000) Analysis of perchlorate in groundwater by
electrospray ionization mass spectrometry/mass spectrometry. Environ Sci Technol 34: 1862-
1864.
Lane, D.J. (1991) 16S/23S rRNA sequencing. In Modern microbiological methods.
Stackebrandt, E., and Goodfellow, M. (eds). Chichester ; New York: Wiley.
Logan, B.E. (1998) A review of chlorate- and perchlorate-respiring microorganisms. Bioremed J
2: 69-79.
24 Logan, B.E. (2001) Assessing the outlook for perchlorate remediation. Environ Sci Technol 35:
482a-487a.
Logan, B.E., and LaPoint, D. (2002) Treatment of perchlorate- and nitrate-contaminated
groundwater in an autotrophic, gas phase, packed-bed bioreactor. Water Res In press.
Logan, B.E., Kim, K., and Price, S. (2001a) Perchlorate degradation in bench- and pilot scale ex-
situ bioreactors. In Bioremediation of Inorganic Compounds. Leeson, A., Peyton, B.M.,
Means, J.L., and Magar, V.S. (eds). Columbus, OH: Battelle Press, pp. 303-308.
Logan, B.E., Zhang, H., Mulvaney, P., Milner, M.G., Head, I.M., and Unz, R.F. (2001b) Kinetics
of perchlorate- and chlorate-respiring bacteria. Appl Environ Microbiol 67: 2499-2506.
Stackebrandt, E., and Gobel, B.M. (1994) Taxonomic note: A place for DNA-DNA reassociation
and 16S rRNA sequence analysis in the present species definition in bacteriology. Int J Syst
Bacteriol 44: 846-849.
Maymo-Gatell, X., Chien, Y.T., Gossett, J.M., and Zinder, S.H. (1997) Isolation of a bacterium
that reductively dechlorinates tetrachloroethene to ethene. Science 276: 1568-1571.
Miller, J.P., and Logan, B.E. (2000) Sustained perchlorate degradation in an autotrophic, gas-
phase, packed-bed bioreactor. Environ Sci Technol 34: 3018-3022.
Nerenberg, R., and Rittmann, B.E. (2002) Perchlorate as a secondary substrate in a denitrifying
hollow-fiber membrane biofilm reactor. Water Sci Technol In press.
Pella, E. (1990) Elemental Organic-Analysis .1. Historical Developments. Am Lab 22: 116-125.
Schilt, A.A. (1979) Perchloric acid and perchlorates. Columbus, Ohio: G. F. Smith Chemical
Co.
Stouthamer, A.H., and Kooijman, S.A. (1993) Why it pays for bacteria to delete disused DNA
and to maintain megaplasmids. Antonie Van Leeuwenhoek 63: 39-43.
25 Thompson, J.D., Gibson, T.J., Plewniak, F., Jeanmougin, F., and Higgins, D.G. (1997) The
CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided
by quality analysis tools. Nucleic Acids Res 25: 4876-4882. van Ginkel, C.G., Rikken, G.B., Kroon, A.G.M., and Kengen, S.W.M. (1996) Purification and
characterization of chlorite dismutase: A novel oxygen-generating enzyme. Arch Microbiol
166: 321-326. van Ginkel, C.G., Gijsbertus., C., Kroon., Maria., A.G., Wijk., V., and Jan., R. (1998) Process
for the degradation of chlorite. United States Patent. 5,891,339.
Wallace, W., Ward, T., Breen, A., and Attaway, H. (1996) Identification of an anaerobic
bacterium which reduces perchlorate and chlorate as Wolinella succinogenes. J Ind Microbiol
16: 68-72.
Wu, J., Unz, R.F., and Logan, B.E. (2001) Persistence of perchlorate and the relative numbers of
perchlorate- and chlorate-respiring microorganisms in natural waters, soils and wastewater.
Bioremed J 5: 119-130.
26 6
5
4
3
2 Perchlorate (mM) Perchlorate
1
0
0 1020304050 Time (d)
Figure 2.1 Perchlorate reduction by the chemolithoautotrophic hydrogen-oxidizing enrichment culture. Arrows indicate culture transfer to fresh media.
Figure 2.2 Scanning electron micrograph of anaerobically grown cells of strain HZ
27 0.4 12 0.3 10 0.2
8
0.1 0.09 6 0.08 0.07 0.06 Chloride (mM) OD (at 600 nm) 4 0.05 0.04 2 0.03
0.02 0 0 122436486072
Time (h)
Figure 2.3 Growth of strain HZ with hydrogen as the electron donor and perchlorate (10 mM) as the electron acceptor with carbon dioxide as the only carbon source. Filled circles (●), cell density in the presence of hydrogen; open circles (○), cell density in the absence of hydrogen; filled squares (■), chloride concentrations in the presence of hydrogen; open squares (□), chloride concentrations in the absence of hydrogen. Error bars (± SD) based on triplicate samples are sometimes smaller than symbol size.
28
Figure 2.4 Neighbor-joining phylogenetic tree of 16S rDNA sequences of strain HZ and others. The same topology was obtained with parsimony and maximum-likelihood analyses. Bootstrap values (100 replicates) are indicated on the nodes. Phylogenetic analysis was based on 1202 nucleotide positions corresponding to Escherichia coli rRNA positions 190-1392.
29 Chapter 3
Molecular assessment of inoculated and indigenous bacteria in biofilms from a pilot-scale perchlorate-reducing bioreactor
Abstract
Bioremediation of perchlorate-contaminated groundwater can occur via bacterial reduction of perchlorate to chloride. Although perchlorate reduction has been demonstrated in bacterial pure cultures, little is known about the efficacy of using perchlorate-reducing bacteria as inoculants for bioremediation in the field. A pilot-scale, fixed-bed bioreactor containing plastic support media was used to treat perchlorate-contaminated groundwater at a site in
Southern California. The bioreactor was inoculated with a field-grown suspension of the perchlorate-respiring bacterium, Dechlorosoma sp. strain KJ, and fed groundwater containing indigenous bacteria and a carbon source amendment. Because the reactor was flushed weekly to remove accumulated biomass, only bacteria capable of growing in biofilms in the reactor were expected to survive. After 26 days of operation, perchlorate was not detected in bioreactor effluent. Perchlorate remained undetected by ion chromatography (detection limit 4 µg L-1) during six months’ operation, after which the reactor was drained. Plastic media was subsampled from top, middle, and bottom locations of the reactor for shipment on blue ice and storage at -
80°C prior to analysis. Microbial community DNA was extracted from successive washes of thawed biofilm material for PCR-based community profiling by 16S-23S ribosomal intergenic spacer analysis (RISA). No DNA sequences characteristic of strain KJ were recovered from any
RISA bands. The most intense bands yielded DNA sequences with high similarities to
Dechloromonas spp., a closely related but different genus of perchlorate-respiring bacteria.
30 Additional sequences from RISA profiles indicated presence of representatives of the low G+C gram-positive bacteria and the Cytophaga-Flavobacterium-Bacteroides group. Confocal scanning laser microscopy and fluorescence in situ hybridization (FISH) were also used to examine biofilms using genus-specific 16S ribosomal RNA probes. FISH was more sensitive than RISA profiling in detecting possible survivors from the initial inoculum. FISH revealed that bacteria hybridizing to Dechlorosoma probes constituted less than 1% of all cells in the biofilms examined, except in the deepest portions where they represented 3-5%. Numbers of bacteria hybridizing to Dechloromonas probes decreased as biofilm depth increased, and they were most abundant at the biofilm surface (23% of all cells). These spatial distribution differences suggested persistence of low numbers of the inoculated strain Dechlorosoma sp. KJ in parts of the biofilm nearest to the plastic medium, concomitant with active colonization or growth by indigenous Dechloromonas spp. in the biofilm exterior. This study demonstrated the feasibility of post hoc analysis of frozen biofilms following completion of field remediation studies.
3.1 Introduction
For decades, discharges of perchlorate-containing wastes were not regulated during the manufacture and disposal of large quantities of ammonium perchlorate, primarily used as oxidizers in rocket propellants and munitions (Espenson 2000; Logan 2001). As many as 75 perchlorate release sites in 22 states have been identified in the U.S. as potential sources of perchlorate contamination of ground and surface waters (Renner 2003). The only known natural source of concentrated perchlorate salts are Chilean nitrate deposits which, when used in agricultural fertilizers, might contribute to some low-level perchlorate dissemination (Schilt
1979). Other recently identified sources of perchlorate include natural evaporites and
31 electrochemical reactions occurring in storage tanks and pipelines. The latter reactions were implicated when high perchlorate concentrations were detected in a water storage tank in
Levelland, Texas (Christen 2003). It is now recognized that perchlorate contamination of subsurface waters at low parts-per-billion concentrations may be much more widespread than previously thought.
Perchlorate is highly soluble, stable, and mobile in aqueous systems (Espenson 2000).
Recent studies have raised concerns that perchlorate, although used as a drug to treat thyrotoxicosis (Wolff 1998), might also inhibit the production of thyroid hormones in healthy populations. Based upon recent epidemiological and toxicological studies, the U. S.
Environmental Protection Agency's January 2002 draft report recommended a reference dose of
0.00003 mg kg-1 day-1. This dose would result in a drinking water standard of 1 µg L-1, using the
assumed factors of 70 kg body weight and 2 L of water consumption per day (Renner 2003). The
California Department of Health Services has responded to the draft report by reducing their
Action Level for perchlorate from 18 µg L-1 to 4 µg L-1, a level very close to the reported
perchlorate detection limit (2.5 to 4 µg L-1) using chromatography-based methods.
Abiotic reduction of perchlorate can be very slow as a result of the high activation energy
required for this process (Gu et al. 2003). However, bacteria capable of reducing perchlorate
during anaerobic respiration are widespread in nature (Coates et al. 1999; Nozawa-Inoue et al.
2003; Wu J 2001), and perchlorate toxicity in bacteria has been observed only at very high
concentrations of 5 g L-1 (Schilt 1979). Perchlorate-reducing bacteria have been isolated from
numerous habitats presumably never exposed to perchlorate (although more comprehensive
testing may lead to a reassessment of extant environmental perchlorate concentrations). A variety
32 of laboratory-isolated pure cultures can use diverse electron donors for complete reduction of perchlorate to chloride via
- - - - ClO4 ClO3 ClO2 Cl + O2 (Rikken et al. 1996)
The first two reactions are catalyzed by dissimilatory perchlorate and chlorate reductases
(Logan et al. 2001b), while the third reaction is catalyzed by the non-respiratory enzyme chlorite dismutase, induced during perchlorate reduction (Chaudhuri et al. 2002; Coates et al., 1999).
Oxygen produced by the third reaction is apparently rapidly consumed, because oxygen concentrations as low as 2 mg L-1 can inhibit perchlorate reduction (Chaudhuri et al. 2002; Song
and Logan 2004). Perchlorate-reducing bacteria are either facultative anaerobes or
microaerophiles that use oxygen preferentially as electron acceptor (Bruce et al. 1999; Coates et
al. 2001; Coates et al., 1999; Herman and Frankenberger 1999; Logan et al. 2001a; Rikken et al.
1996, Wallace et al. 1996; Wu J 2001; Zhang et al. 2002). Many but not all perchlorate-reducing
bacteria are also capable of dissimilatory nitrate reduction, suggesting that the two pathways are
separate (Bruce et al. 1999; Coates et al., 1999).
The β-subdivision of Proteobacteria contains two groups of closely related perchlorate
degraders, Dechloromonas spp. and Dechlorosoma spp., which are particularly widespread in the
environment (Achenbach et al. 2001). The most thoroughly characterized representatives of
these two groups are Dechloromonas agitata and Dechlorosoma suillum, respectively. Although
D. suillum is now recognized as Azospira suillum (Tan and Reinhold-Hurek 2003), we refer to
the latter group of perchlorate degraders in this report as Dechlorosoma spp. Both groups include
representatives that can reduce both perchlorate and nitrate (Chaudhuri et al. 2002). One such
strain, Dechlorosoma sp. KJ, was inoculated into a pilot-scale packed-bed bioreactor at a site in
California for ex situ treatment of groundwater contaminated with both perchlorate and nitrate
33 (Min et al. 2003). Although other engineered bioreactor systems have also been designed and used to treat perchlorate-contaminated water (Brown et al. 2002; Giblin et al. 2000; Nerenberg et al. 2000; Wallace et al. 1998), little is known about how well the bioaugmented microbes can compete with indigenous populations from the contaminated sites. Interestingly, in a study of trichloroethylene (TCE) degradation using microcosms inoculated with Burkholderia sp., it was concluded that TCE degradation was due not to the inoculated cultures, but to activity by indigenous populations selected by the operating conditions in the microcosms (Munakata-Marr et al. 1997).
The objective of the current study was to analyze biofilm material from frozen samples of reactor packing to determine whether the inoculated strain, Dechlorosoma sp. KJ, could be detected after six months of bioreactor operation. To this end we extracted biofilm community
DNA for RISA (Ribosomal Intergenic Spacer Analysis) (Bormann and Triplett 1997) and examined biofilm material using fluorescent in situ hybridization (FISH) with oligonucleotide probes specific for Dechlorosoma and Dechloromonas spp. We hypothesized that the indigenous microorganisms from groundwater had colonized the bioreactor and outcompeted the inoculated strain.
3.2 Materials and Methods
Bioreactor inoculum. Dechlorosoma sp. strain KJ was originally isolated from a bench- scale, perchlorate-reducing bioreactor (Logan et al. 2001a). This isolate was subsequently shown to reduce nitrate and chlorate as well as perchlorate, but not sulfate (Kim and Logan 2001; Logan et al. 2001a). Because KJ could also reduce nitrate, it was selected as the inoculum strain to treat groundwater containing perchlorate as well as nitrate. Prior to inoculation, KJ was grown for one
34 week in a nitrogen-gas-purged 208-L drum containing groundwater amended with 300 mg L-1
-1 NaClO4, 450 mg L NaCH3COO, and a combined phosphate buffer-nitrogen solution of 1.92 g
-1 K2HPO4, 0.98 g NaH2PO4H2O, and 0.5 g NH4H2PO4 L (Min et al. 2003). Perchlorate
degradation was confirmed in the drum, and the cell suspension amended once more with
perchlorate and nutrients before being pumped into the reactor, which was then left undisturbed
for one day to allow bacteria to attach to media surfaces. The reactor was operated for 11 days in
full recirculation mode using the 208-L drum and then switched to continuous-feed mode for
groundwater treatment.
Bioreactor operation and sampling. Bioreactor dimensions were 2.1 m in height, 0.61 m
in width, and 0.30 m in length. The bioreactor compartment was packed to a height of 1.2 m with
plastic media (Tri-pack, Jaeger Products, Inc.) having 3.175-cm diameter and specific surface
area of 230 m2 m-3 (Min et al. 2003). A perforated plate was placed on top of the plastic media
bed to keep it in the reactor compartment because the density of the plastic was less than that of
water. Groundwater stored in an equalization tank was pumped in upflow mode into the bottom of the bioreactor. At the top of the reactor, water overflowed a weir for transport to a drainage
system by gravity (Min et al. 2003). Two different hydraulic loading rates of 0.34 and 0.68 L m-2
s-1 were tested. These rates corresponded to flows of 3.8 and 7.6 L min-1 and hydraulic detention
times of 56 min and 28 min, respectively. Perchlorate was undetectable in bioreactor effluent at
a flow rate of 0.34 L m-2 h-1, but it was detectable at 0.68 L m-2 h-1, as shown by the intra-column
perchlorate profiles in Figure 3.1. After 26 days of operation at the lower rate, no perchlorate
was detected in bioreactor effluent and it remained undetectable throughout the remainder of the
test period. To remove excess biomass and prevent clogging or short circuiting, the reactor was
35 also flushed from the bottom on a weekly basis by simultaneously increasing groundwater flow and aerating with a compressed air tank (2.6 x 10-3 m3 s-1) for 10 min.
The reactor was fed groundwater containing perchlorate (50-120 µg L-1), dissolved
oxygen (8 – 10 mg L-1), nitrate (4 – 4.5 mg L-1 as N), sulfate (33 mg L-1), as well as added acetic
acid (50 mg L-1) and ammonium phosphate (12.8 mg L-1). No effort was made in this study to
optimize nutrient concentrations, and acetic acid and nitrogen-phosphate solutions (average C:N
molar ratio of 5:1) were added in excess to avoid nutrient limitation. After six months of
operation, flow was turned off and the water drained from the reactor compartment. Individual
pieces of the plastic medium were collected aseptically from the top, middle and bottom of the
reactor. Samples were placed in separate Ziploc bags, shipped to Penn State overnight on blue
ice, and stored at -80oC for one year until analysis.
Water testing. Nine taps were located along one side wall of the compartment to permit
water sampling at different heights. Perchlorate, acetate, nitrate, and sulfate were measured using
ion chromatography (Dionex Corp.) as previously described (Min and Logan 2003). DO
(minimum detection level of 0.3 mg L-1) and oxidation-reduction potential (ORP) were measured
using field instruments (YSI 600 XL). The average influent perchlorate concentration in the
groundwater over 185 days of operation was 75 + 13 µg L-1 (range 59-118 µg L-1; n = 94). The
bioreactor did not completely remove acetic acid, the concentration of which averaged 20 + 8 mg
L-1 at the outlet (Figure 3.2). Intra-column profiles indicated sequential removal of dissolved
oxygen, nitrate, and perchlorate (to less than 1 mg L-1), but no sulfate removal was observed.
Both oxygen and nitrate were undetectable after the water had reached a distance of 0.56 m from
the inlet. Perchlorate was undetectable at 0.85 m. Influent and effluent DO means and standard
36 deviations were 8.7+0.4 and 0.2+0.3 mg L-1, respectively (Figure 3.2). Influent and effluent ORP means and standard deviations were 8+52 and -85+77 mV, respectively.
DNA extraction, PCR, cloning, and 16S rRNA gene sequencing. To obtain biomass
samples, 3-4 pieces of plastic medium were aseptically cut to fit into a 50-ml Falcon tube with 8
ml Tris-EDTA buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) and vortexed at high speed for 1
min. The buffer was decanted and centrifuged to recover cell material from this “first wash.” The
same plastic pieces were rinsed with fresh buffer and drained before another 8 ml buffer was
added and the sample was vortexed and decanted a second time. This wash procedure was
repeated a third time. Biomass samples were designated by their location in the bioreactor (top,
T; middle, M; bottom, B) and by the number of washes prior to their recovery (1, 2, 3). Cell pellets were obtained from wash suspensions by centrifugation. Microbial community DNA was
extracted from the cell pellets by a modified phenol-chloroform method (Bruns and Buckley
2001). The quantity and quality of DNA in extracts was evaluated by absorbances at 260 and 280
nm using a UV spectrophotometer as well as by inspection of stained bands after electrophoresis
in 0.7 % (w/v) agarose gels (Sambrook et al. 1989).
Bacterial community DNA (10 ng) was added as template in polymerase chain reactions
(PCR) using a universal bacterial primer set which amplifies the 3’-ends of 16S rRNA genes,
along with the 16S-23S intergenic spacer regions and short 5’-end pieces of 23S rRNA genes
(Bruns et al. 2001). Because the lengths of bacterial spacers vary over a range of 80-1500 bp, the
amplification products can be separated by size through gel electrophoresis to obtain bacterial
community profiles (Bormann and Triplett 1997). Individual DNA bands can be excised from
the gels and cloned to obtain sequences (ca. 600 bp) from 3’-portions of the 16S rRNA genes.
We used forward and reverse primers 926F (5’-AAACTYAAAKGAATTGACGG-3’) and
37 23S115R (5’-GGGTTBCCCCATTCGG-3’), respectively (Lane et al. 1991). The PCR products were separated by electrophoresis (80 Volts for 16 hours) in 8 % polyacrylamide gels, which were stained with SYBR-Green (Molecular Probes) for visualization by UV illumination. Gel bands were excised with a razor blade, and DNA was eluted from the bands, extracted and precipitated by using standard methods (Sambrook et al. 1989). The DNA was ligated into a plasmid vector (TOPO TA, Invitrogen) for cloning into competent E. coli cells. DNA sequences were obtained from at least two purified transformants for each band (ABI Hitachi 3100).
Sequence editing and alignment were conducted by using EditSeq and Seqman programs
(Lasergene). The CHIMERA_CHECK (Cole et al. 2003) and Bellerophon (Hugenholtz and
Huber 2003) programs were used to evaluate sequences for chimeras. Closest relatives of the retrieved sequences were identified by searching GenBank with the Basic Local Alignment
Search Tool (Altschul et al. 1990).
Preparing samples for in situ hybridization. Biofilm samples for hybridization and microscopic examination were obtained from thawed pieces of the plastic medium. A clean, ethanol-wiped scalpel was used to scrape off the surface layer of the plastic medium along with the biofilm in order to examine the entire depth of the biofilm on the plastic surface. The excised pieces (each approximately 6 mm by 6 mm) were placed on MultiwellTM coverslips (Molecular
Probes) with the outer side of the biofilm facing the coverslip. In order to keep the biofilms from
dislodging during the hybridization and washing steps, they were embedded in 10 µl 0.7 %
agarose (Moller et al. 1996). Sample fixation, washing, and dehydration were performed
following standard procedures (Manz et al. 1999).
Oligonucleotide probes. The probe sequences, their complementary sites on 16S rRNA
genes, and the conditions used in hybridization and washing are summarized in Table 1. The
38 sequence of Dechloromonas probe Monas1403 was identified by a manual alignment with all other Dechloromonas and Dechlorosoma 16S rRNA sequences in Genbank. The Dechlorosoma probe Soma1035 was originally used as a PCR primer (data not shown). The specificity of the probes was evaluated by BLAST against the Genbank database, as well as by the
CHECK_PROBE program from the Ribosomal Database Project II (Cole et al. 2003). The 5’ end labels for probes EUB338, Monas1403, and Soma1035 were Cy5, Cy3, and FITC, respectively.
To confirm probe specificity, cell suspensions from pure cultures of Dechloromonas sp. HZ and
Dechlorosoma sp. KJ were used as positive and negative controls. A nonsense probe,
NONEUB338 whose nucleotide sequence complementary to that of probe EUB338, was used as a control for nonspecific hybridization. Cell suspensions were prepared by fixation with paraformaldehyde at a final concentration of 1 % for 90 minutes, washing with phosphate- buffered saline (PBS, 130 mM sodium chloride, 10 mM sodium phosphate buffer at pH 7.2), resuspension in 1:1 mixture of PBS:96% ethanol and storage at -20 oC (Amann et al. 1990).
Fluorescent in situ hybridization. For cell suspensions, FISH was performed according
to established procedures (Amann et al. 1990; Amann et al. 1991). Briefly, cells were spotted
onto heavy Teflon coated slides (Erie Scientific Company), air dried, dehydrated in an ethanol
gradient, hybridized with oligonucleotide probes under various amount of formamide, and
washed with buffer containing appropriate amounts of sodium chloride (Table 1). For biofilm
samples, hybridization was done at 46 oC for 16 h in a moist chamber and washed at 48 oC for 1 h as suggested by Moller and coworkers (Moller et al. 1996).
Image acquisition and quantitation of biofilm coverage on the support surface. Two separate biofilm samples from the top location were selected for finer-scale analysis by confocal laser scanning microscopy. To obtain images of cells stained with probes labeled with Cy5, Cy3,
39 and FITC (fluorescein isothiocyanate), three excitation laser lines, red (633 nm), green (543 nm), and blue (488 nm), respectively, were used with an Olympus Fluoview 300 confocal microscope to acquire images in three independent channels. Images were viewed with a Plan-Apo 60X oil objective (numerical aperture = 1.4). The z direction images were acquired at 3-µm intervals down into the biofilm. The 16-bit gray scale images from each channel were first converted to 8- bit gray scale images required for COMSTAT input (Heydorn et al. 1995). A fixed threshold value was used to generate binary images for each channel based on the ability to distinguish cells from background. The threshold gray values for Cy5, Cy3, and FITC channels were 80, 35, and 17, respectively. Images were examined manually at different depths throughout the z stacks, and no appreciable quenching of fluorescent signal was observed within the depth range used.
Threshold-processed images were then subjected to COMSTAT analysis. Relative abundance of cells hybridizing with the Monas1403 and Soma1035 probes were determined by using the function of “area occupied by bacteria in each layer” in the COMSTAT program (Heydorn et al.
1995).
For the determination of biofilm thickness, agarose-embedded samples were stained with the BacLight LIVE/DEAD viability dyes (Molecular Probes Inc., Eugene, OR). When stained, the biofilm matrix exhibited a uniform but low background fluorescence that was readily distinguishable from the brightly fluorescing, intact cells. The agarose exhibited very little fluorescence. The thickness of the biofilm was determined with an UplanFL 40X objective
(numerical aperture = 0.75) by moving the focal plane away from the plastic surface until no fluorescence was observed. The distance between initial and final positions of the objective lens was used to calculate biofilm thickness. Estimates of mean biofilm thickness from top, middle, and bottom locations of the bioreactor were obtained from 3 or 4 independent measurements for
40 each location. Total scanned areas for mean thickness calculations for top, middle, and bottom samples were 3.75 x 105, 3.74 x 105, and 4.99 x 105 µm2, respectively.
GenBank accession numbers for partial 16S rRNA genes (E. coli positions 907-1542) are
AY515710-AY515723.
3.3 Results
DNA was recovered from all three washes of plastic media from the top and middle
locations, but only the first wash from the bottom location yielded enough DNA for PCR amplification. RISA community profiles from biofilm material removed by successive washes were compared to the RISA profile from genomic DNA of a pure culture of Dechlorosoma sp. strain KJ (Figure 3.3). None of the biofilm community profiles from the top location contained a band that migrated to the same position in the gel as the single RISA band from the
Dechlorosoma sp. strain KJ culture. A putative KJ band could not be ruled out by visual inspection of the community profiles from the middle and bottom locations. Bands at positions nearest to that of KJ, however, yielded DNA sequences most closely related to representatives of the Cytophaga-Flavobacterium-Bacteroides (CFB) group and low G+C gram-positive bacteria
(Table 2). Although no sequences from Dechlorosoma sp. KJ were recovered from any RISA profile bands, biofilms from all bioreactor locations contained intense bands yielding DNA sequences from Dechloromonas spp., the other genus within the β-Proteobacteria which is known to reduce perchlorate (1). Except for T1-3, which is a chimera detected by the
Bellepheron program, all other sequences fell within the CFB and low G+C gram-positive groups (Table 2).
41 Biofilms from the top and middle locations of the bioreactor were thicker than biofilms from the bottom location, which presumably reflected the effect of upward flushing from the reactor bottom. Measured thicknesses of biofilms from the top and middle locations were
96.7±25.5 and 89.3±12.9 µm (mean ± standard deviation, n = 3), respectively. The thickness of the biofilm from the bottom-location was 57.5±22.5 µm (n = 4). Community profiles from the first and second washes of plastic packing from the top location did not contain the same bands as the community profile from the third wash, indicating qualitative differences in populations that could be dislodged with different degrees of washing force. The three bands having different positions in the T3 community profile all yielded DNA sequences belonging to low G+C gram- positive bacteria (Table 2).
To assess presence and spatial distribution of Dechlorosoma and Dechloromonas spp. on a finer scale, we applied FISH probes Soma1035 and Monas1403, respectively, in conjunction with a universal EUB338 probe, to biofilm samples for analysis with confocal microscopy. Probe specificities were validated in hybridization tests with cells from pure cultures of the respective target organisms (Figures 3.4A and 3.4B), and no cross hybridization was observed (data not shown). Nonspecific hybridization to probe NONEUB338 also was not observed. Confocal microscopic images of probe-hybridized biofilm samples are shown in Figure 3.4C. Sequential analysis of Z stack images indicated that most cells hybridizing to the Monas1403 probe (pink color in Figure 3.4C) were in the exterior portions of the biofilm (0 and 15 µm depths). Similar distribution patterns were observed in three independently selected fields-of-view. Very low numbers of cells hybridized with the Soma1305 probe. These were indicated by signals from the green channel, which appeared cyan in combination with the blue-channel signal from co- hybridization with EUB338 (arrows in last panel of Figure 3.4C).
42 Relative abundances of bacteria hybridizing to the Monas and Soma probes were determined on the basis of total field-of-view areas using the COMSTAT program (Figure 3.5A).
Only 50% of total field area was covered by cells hybridizing to the EUB338 probe, with the remainder of the field area exhibiting a uniform background fluorescence. Cells hybridizing to the Monas and Soma probes covered 12% and <1% of total field areas, respectively. Relative abundances of Dechloromonas and Dechlorosoma spp. were also assessed in relation to total cells hybridizing with probe EUB338 (Figure 3.5B). The results clearly showed that abundance of cells hybridizing with the Monas probe (Dechloromonas spp.) decreased with depth in the biofilm. While Dechloromonas spp. accounted for nearly 23% of bacterial biomass at the surface
(z = 0 µm), this proportion decreased to only about 1% at a depth of 111 µm from the surface.
Cells hybridizing with the Soma probe (Dechlorosoma spp.) represented less than 1% of bacterial biomass throughout most of the biofilm. At the bottom of the biofilm near the plastic surface, however, Dechlorosoma spp. constituted about 5% of all bacterial biomass (Figure
3.5B), suggesting that these may have been surviving populations from the original inoculum.
3.4 Discussion
After six months of operation following inoculation with Dechlorosoma sp. KJ, the perchlorate-reducing bioreactor supported biofilms comprising multiple species representing three phylogenetic groups: β-Proteobacteria, the CFB group, and low G+C gram-positive bacteria. Biofilm populations derived from the Dechlorosoma sp. KJ inoculum were not in sufficiently high numbers to be detected by PCR-based community profiling. Instead, RISA detected greater abundances of members of the genus Dechloromonas spp., which are also known to reduce perchlorate. DNA sequences characteristic of Dechloromonas spp. were
43 recovered from prominent bands in community profiles from all three locations of the bioreactor.
Lack of detection by RISA of DNA sequences for Dechlorosoma spp. may have been due to the fact that KJ was inoculated only once into the bioreactor. Since the reactor was also flushed on a weekly basis to remove excess buildup of biomass, regular disturbance may have interfered with the ability of inoculated populations to persist (Min and Logan 2003). Weekly flushing to remove excess biomass may also have favored the establishment of indigenous bacteria that were continually being reintroduced with the groundwater being treated.
Another reason why KJ did not become dominant in the biofilms could be the relatively low concentration of perchlorate in the groundwater feed (0.05-0.12 mg L-1), compared to the
300 mg L-1 perchlorate in the medium in which KJ had been grown and acclimated prior to
inoculation of the bioreactor (Min and Logan 2003). It should also be noted that KJ was
originally isolated from a laboratory reactor fed with medium containing 20 mg L-1 perchlorate
(Kim and Logan 2001) and subsequently determined to have a half saturation constant for
perchlorate of 33 mg L-1 (Logan et al. 2001a). Indigenous bacteria in the groundwater could
therefore have been adapted better to the bioreactor’s perchlorate concentrations. Furthermore,
perchlorate represented such a small proportion of the total electron acceptor pool that a very
small population would have been sufficient to reduce it. Microbial populations, particularly
those at the bottom of the reactor, were exposed to influent concentrations of dissolved oxygen
and nitrate that were 80 fold higher than that of perchlorate. The nutrient and operating
conditions of the bioreactor would therefore not have enriched solely for perchlorate-reducing
bacteria. There was also a significant amount of residual acetic acid in the reactor effluent (21
mg L-1), indicating that electron donor availability was ample for diverse populations to become
established.
44 In considering solely the perchlorate-reducing populations, the presence of dissolved oxygen and nitrate in the groundwater may also have selected for perchorate reducers that were less sensitive to fluxes in electron acceptors. In a study by Chaudhuri et al., Dechlorosoma suillum and Dechloromonas agitata strain CKB exhibited clear differences in activity when transferred from media containing only one type of electron acceptor (perchlorate or nitrate) to media containing both (Chaudhuri et al. 2002).When transferred to media containing equimolar amounts of perchlorate and nitrate, nitrate-grown cells of D. suillum exhibited an extended lag, while perchlorate-grown cells preferentially reduced nitrate and did not reduce perchlorate until nitrate was removed completely. In contrast, D. agitata strain CKB reduced perchlorate equally well in the presence or absence of nitrate (Chaudhuri et al. 2002). In other studies conducted on
Dechlorosoma sp. KJ, chlorate and nitrate reduction pathways were observed to be induced
- -1 separately, and the presence of as little as 5 µg NO3 L inhibited its ability to reduce chlorate
(Xu et al. 2004). These studies are consistent with our observations in the nitrate- and
perchlorate-fed bioreactor, where cells hybridizing with the Monas1403 probe outnumbered cells
hybridizing with the Soma1035 probe. The ability to simultaneously reduce nitrate and
perchlorate would have been a selective advantage for populations in our bioreactor. Before this
could be considered a definitive explanation for our results, however, other strains of
Dechloromonas spp. would need to be evaluated for their ability to carry out simultaneous
reduction of nitrate and perchlorate.
In our study, FISH probes could have hybridized to some beta-proteobacteria belonging
to genera other than Dechlorosoma and Dechloromonas spp. RDP II check_probe results for
Monas1403 yielded a total of 17 perfect matches consisting of 8 sequences from Dechloromonas
spp., and 9 sequences from other beta-Proteobacteria including 2 Leptothrix, 1 Acidovorax, and 6
45 unaligned sequences. Results for Soma1305 yielded 7 matches, all within the Rhodocyclus tenuis subgroup (4 Dechlorosoma spp., 2 Azoarcus, and 1 uncultured clone). Evidence in our study which supports FISH detection of Dechloromonas populations, however, is the independent recovery of 600-bp rRNA gene sequences representative of Dechloromonas spp. from RISA profiles.
Based on the reactor chemical profiles in Figure 3.2, DO and nitrate concentrations at the bottom location would not have enabled induction of perchlorate reduction by Dechlorosoma sp.
KJ. Although the depletion of DO and nitrate at the middle and top locations would have been more permissive, rRNA gene sequences associated with the inoculum strain were not detected in
RISA profiles at these locations. As expected, FISH and image analysis at a finer spatial scale were more sensitive in detecting less abundant populations. FISH showed that bacteria hybridizing to the Dechlorosoma probe were present in very low numbers in the deep interior of top-location biofilms (111 µm from the biofilm surface), although they were outnumbered by
Dechloromonas spp. at more shallow depths (Figures 3.5A and 3.5B). The numbers of cells hybridizing with the Dechloromonas probe increased with distance from the plastic medium, reaching their highest proportions of total bacterial biomass at the surface. These spatial differences suggested that Dechloromonas populations were introduced from groundwater.
Although the inoculated Dechlorosoma sp. strain KJ did not maintain a dominant presence in biofilms, the bioreactor was still effective at removing perchlorate, presumably due to the establishment and activity of perchlorate-reducing, indigenous populations. Our results are not the first to show that indigenous microorganisms can outcompete bioaugmented bacteria derived from laboratory strains. In one study of bioreactors treating mercury-contaminated wastes, two mercury-resistant inoculum strains disappeared after 485 days, whereas two different
46 γ-proteobacteria capable of reducing mercury invaded the biofilms (Canstein et al. 2001). In a microcosm study of TCE degradation, indigenous bacteria rather than inoculated strains were hypothesized to be responsible for TCE degradation (Munakata-Marr et al. 1997).
Despite a long-held recognition that microorganisms cultured from environmental samples are often not the most dominant or active populations in situ, some laboratory enrichment and isolation procedures have nevertheless yielded closely related representatives of previously uncharacterized and important indigenous bacteria. In a controlled field study of in situ MTBE degradation in groundwater, for example, ribosomal RNA gene sequences with high similarity to laboratory strain PM1 were detected in higher numbers following a rise in MTBE concentration (Hristova et al. 2003). We present here another example of independent molecular detection of members of a recently recognized genus, Dechloromonas, which was identified through traditional cultural procedures and has demonstrated capabilities for degrading a specific pollutant (Achenbach et al. 2001; Coates et al., 1999; Kim and Logan 2001; Logan et al. 2001a).
Representatives of the other phylogenetic groups detected in the biofilms are not known to have the ability to reduce perchlorate (Achenbach et al. 2001; Logan 2001b). Although their roles are unknown, members of the CFB group are widespread and known to utilize diverse carbon substrates (Kirchman 2002). The rRNA gene sequences belonging to the CFB group in our bioreactor had high similarities to GenBank accessions from diverse environments, including deep well groundwater at a site in Russia (clones from T1-1 and B1-1), and stable bacterial consortia for chlorobenzene treatment (clone from T2-5). The five rRNA gene sequences recovered from our reactor that belonged to low G+C gram-positive bacteria were genetically less diverse than the CFB sequences. Three of the five clones (from T1-2, T3-1, and M3-4) had identical 16S rRNA gene sequences (600 nt at 3’-ends) with greatest similarity (90%) to a
47 GenBank accession from coal-tar-waste-contaminated aquifer waters (Bakermans et al. 2002).
The other two rRNA gene sequences (from T3-3 and T3-4) both had highest similarities (99%) to a GenBank accession from benzene-contaminated groundwater. The recovery of nearly identical rRNA gene sequences from bands in different locations of the RISA gels indicated that some biofilm populations, in particular gram-positive bacteria, could have had more than one ribosomal RNA operon.
In this study we followed a stepwise molecular approach to analyzing biofilm communities in the perchlorate-degrading bioreactor. In the initial, exploratory step, we used
RISA profiling with universal bacterial primers to identify dominant populations and discern qualitative differences based on bioreactor location and ease of removal from the plastic medium. RISA analysis enabled us to identify bacterial populations other than the inoculated strain which appeared to be dominant in biofilm communities. Secondly, cloning and sequencing of the 16S rRNA gene portions of excised RISA bands provided the basis for selecting the most informative probes for key populations. Third, we tried FISH analysis to obtain finer scale resolution and more quantitative estimates of biofilm populations, recognizing that freezing and thawing of the biofilm samples could have affected biofilm integrity. Although only a very small fraction of total surface area of plastic media was analyzed in our study, our objective was not to conduct a comprehensive spatial analysis of the bioreactor but to assess the feasibility of post hoc molecular techniques. Our study demonstrated that FISH can be performed successfully on some biofilm materials even after frozen storage and that FISH might be applicable in other studies to relate biofilm community composition to remediation performance. Finally, use of genus-specific probes in FISH analysis permitted quantification of population sizes in relation to
48 biofilm depth and provided the basis for evaluating the utility of bioaugmentation in this pilot- scale system.
Acknowledgements
We thank Arne Heydorn for providing the COMSTAT program and Ian Head for review comments. This research was funded by the American Water Works Association Research
Foundation 2557. H.Z. was supported by the Penn State Biogeochemical Research Initiative for
Education (BRIE) sponsored by NSF (IGERT) grant DGE-9972759.
49 Table 3.1: FISH probe sequences, target sites, formamide concentrations in the hybridization buffer and sodium chloride concentrations in the washing buffer.
Target 16S rRNA % Formamide in NaCl in the wash Probe positions (E. coli Probe sequence Reference hybridization buffer buffer (mM) numbering)
EUB338 338-355 5'-GCTGCCTCCCGTAGGAGT-3’ 20 225 (3)
NONEUB 338-355 5 -ACTCCTACGGGAGGCAGC-3 20 225 (44)
Monas1403 1403-1419 5’-GCGGAACCCGCTCCCAT-3’ 20 225 This study
Soma1035 1017-1035 5'-CCATCTCTGGAAAGTTCCTGG-3’ 0 900 This study
50 Table 3.2 Phylogenetic summary of perchlorate-reducing community from cloning and sequencing results (only 16S rDNA portion up to “GATCAC” were used in BLAST search).
Band and Main Group Closest relatives in GenBank Accession % accession number similarity number
T1-1, B1-1 CFB Clone S15A-MN90 AJ534684 98 (AY510710, (deep well groundwater in Russia) AY515714) T1-2, T3-1, Low G+C gram pos Clone:KU8 (coal-tar-waste- AB074938 90 M3-3 contaminated aquifer water) (AY515711, AY515720, AY515721) T1-4, B1-4 β-Proteobacteria Dechloromonas sp. strain JJ AY032611 98 (AY515718, AY515722) T2-4 β-Proteobacteria Dechloromonas sp. strain HZ AF479766 99 (AY515712) T2-5 CFB Clone IA-5 AJ488076 94 (AY515719) (chlorobenzene-treatment consortia)
T3-3, T3-4 Low G+C gram pos Clone ZZ12C4 (benzene- AY214180 98 (AY515713, contaminated groundwater) AY515715) M1-6 β-Proteobacteria Dechloromonas sp. strain JM AF323489 100 (AY515723) B1-2 CFB Flavobacterium psychrophilum AB078060 97 (AY515716)
51 3.5 References
Achenbach L, Michaelidou U, Bruce R, Fryman J, Coates J (2001) Dechloromonas agitata gen.
nov., sp. nov. and Dechlorosoma suillum gen. nov., sp. nov., two novel environmentally
dominant (per)chlorate-reducing bacteria and their phylogenetic position. Int J Syst Evol
Microbiol 51:527-533
Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search
tool. J Mol Biol 215:403-410
Amann RI, Binder BJ, Olson RJ, Chisholm SW, Devereux R, Stahl DA (1990) Combination of
16S ribosomal-RNA-targeted oligonucleotide probes with flow-cytometry for analyzing
mixed microbial populations. Appl Environ Microbiol 56:1919-1925
Amann RI, Krumholz L, Stahl DA (1990) Fluorescent-oligonucleotide probing of whole cells for
determinative, phylogenetic, and environmental studies in microbiology. J Bacteriol
172:762-770
Bakermans C, Hohnstock-Ashe AM, Padmanabhan S, Padmanabhan P, Madsen EL (2002)
Geochemical and physiological evidence for mixed aerobic and anaerobic field
biodegradation of coal tar waste by subsurface microbial communities. Microb Ecol
44:107-117
Borneman J, Triplett EW (1997) Molecular microbial diversity in soils from eastern Amazonia:
evidence for unusual microorganisms and microbial population shifts associated with
deforestation. Appl Environ Microbiol 63:2647-2653
Brown JC, Snoeyink VL, Kirisits MJ (2002) Abiotic and biotic perchlorate removal in an
activated filter. J Amer Water Works Assoc 94:70-79
52 Bruce RA., Achenbach LA, Coates JD (1999) Reduction of (per)chlorate by a novel organism
isolated from paper mill waste. Environ Microbiol 1:319-329
Bruns MA, Buckley DH (2001) Isolation and purification of microbial community nucleic acid
from environmental samples. In: C Hurst et al (eds), Manual of Environmental
Microbiology, 2nd ed. ASM.Press, Washington, DC, pp 564-572
Bruns MA, Hanson JR, Mefford J, Scow KM (2001) Isolate PM1 populations are dominant and
novel methyl tert-butyl ether-degrading bacterial in compost biofilter enrichments.
Environ Microbiol 3:220-5
Canstein HF, Li Y, Felske A, Wagner-Dobler I (2001) Long-term stability of mercury-reducing
microbial biofilm communities analyzed by 16S-23S rDNA interspacer region
polymorphism. Microb Ecol 42:624-634
Chaudhuri SK, O’Connor SM, Gustavson RL, Achenbach LA, Coates JD (2002) Environmental
factors that control microbial perchlorate reduction. Appl Environ Microbiol 68:4425-
4430.
Christen K (2003) Perchlorate mystery surfaces in Texas. Environ Sci Technol 37:376A-377A
Coates J D, Chakraborty R, Lack JG, O'Connor SM, Cole KA, Bender KS, Achenbach LA
(2001) Anaerobic benzene oxidation coupled to nitrate reduction in pure culture by two
strains of Dechloromonas. Nature 411:1039-1043
Coates JD, Michaelidou U, Bruce RA, O'Connor SM, Crespi JN, Achenbach LA (1999)
Ubiquity and diversity of dissimilatory (per)chlorate-reducing Bacteria. Appl Environ
Microbiol 65:5234-5241
Cole JR, Chai B, Marsh TL, Farris RJ, Wang Q, Kulam SA, Chandra S, McGarrell DM, Schmidt
TM, Garrity GM, Tiedje JM (2003) The Ribosomal Database Project (RDP-II):
53 previewing a new autoaligner that allows regular updates and the new prokaryotic
taxonomy. Nucl Acids. Res 31:442-443
Espenson JH (2000) The problem and perversity of perchlorate. In: ET Urbansky (ed),
Perchlorate in the Environment. Kluwer Academic/Plenum, New York, pp 1-8
Giblin T, Herman D, Deshusses MA, Frankenberger WT (2000) Removal of perchlorate in
ground water with a flow-through bioreactor. J Environ Qual 29:578-583
Gu B., Dong W, Brown GM, Cole DR (2003) Complete degradation of perchlorate in ferric
chloride and hydrochloric acid under controlled temperature and pressure. Environ Sci
Technol 37:2291-2295
Herman DC, Frankenberger WT (1999) Bacterial reduction of perchlorate and nitrate in water. J
Environ Qual 28:1018-1024
Heydorn A, Nielsen AT, Hentzer M, Sternberg C, Givskov M, Ersboll BK, Molin S (2000)
Quantification of biofilm structures by the novel computer program COMSTAT.
Microbiology 146 ( Pt 10):2395-407
Hristova K, Gebreyesus B, Mackay D, Scow KM (2003) Naturally occurring bacteria similar to
the methyl tert-butyl ether (MTBE)-degrading strain PM1 are present in MTBE-
contaminated groundwater. Appl Environ Microbiol 69:2616-2623
Hugenholtz P, Huber T (2003) Chimeric 16S rDNA sequences of diverse origin are
accumulating in the public databases. Int J Syst Evol Microbiol 53 289-293
Kim K, Logan BE (2001) Microbial reduction of perchlorate in pure and mixed culture packed-
bed bioreactors. Water Res 35:3071-3076
Kirchman DL (2002) The ecology of Cytophaga-Flavobacteria in aquatic environments. FEMS
Microbiol Ecol 39:91-100
54 Lane DJ (1991) 16S/23S rRNA sequencing. In: E Stackebrandt and M Goodfellow (eds) Nucleic
Acid Techniques in Bacterial Systematics. John Wiley & Sons, New York, pp 115-175
Logan BE (2001) Assessing the outlook for perchlorate remediation. Environ Sci & Technol
35:482a-487a
Logan BE, Zhang H, Mulvaney P, Milner MG, Head IM, Unz RF (2001) Kinetics of perchlorate-
and chlorate-respiring bacteria. Appl Environ Microbiol 67:2499-2506.
Manz W (1999) In situ analysis of microbial biofilms by rRNA-targeted oligonucleotide probing.
Methods Enzymol 310:79-91
Massol-Deya A, Whallon J, Hickey R, Tiedje JM (1995) Channel structures in aerobic biofilms
of fixed-film reactors treating contaminated groundwater. Appl Environ Microbiol
61:769-777
Min B, Evans PJ, Chu A, Logan BE (2004) Perchlorate removal in sand and plastic media
bioreactors. Water Res 38:47-60
Moller S, Pedersen AR, Poulsen LK, Arvin E, and Molin S (1996) Activity and three-
dimensional distribution of toluene-degrading Pseudomonas putida in a multispecies
biofilm assessed by quantitative in situ hybridization and scanning confocal laser
microscopy. Appl Environ Microbiol 62:4632-4640
Munakata-Marr J, Matheson VG, Forney LJ, Tiedje JM, McCarty PL (1997) Long-term
biodegradation of trichloroethylene influenced by bioaugmentation and dissolved oxygen
in aquifer microcosms. Environ Sci & Technol 31:786-791
Nerenberg R, Rittmann BE (2002) Perchlorate as a secondary substrate in a denitrifying hollow-
fiber membrane biofilm reactor. Water Science and Technology 2:259–265
55 Nozawa-Inoue M, Leong J, Rolston D, Scow KM (2003) Abstracts of the American Society for
Microbiology General Meeting, May 18th to 22nd, Washington, DC
Renner R (2003) Environmental health, Academy to mediate debate over rocket-fuel
contaminants. Science 299:1829
Rikken GB, Kroon AGM, vanGinkel CG (1996) Transformation of (per)chlorate into chloride
by a newly isolated bacterium: Reduction and dismutation. Appl Microbiol and
Biotechnol 45:420-426
Sambrook J, Fritsch EF, Maniatis T (1989) Molecular cloning: a laboratory manual, 2nd ed. Cold
Spring Harbor Laboratory Press, Plainview, NY
Schilt AA (1979) Perchloric Acid and Perchlorates. G. F. Smith Chemical Co., Columbus, Ohio
Song Y, Logan BE (2004) Effect of O2 exposure on perchlorate reduction by Dechlorosoma sp.
KJ. Water Res. 38:1626-1632
Tan Z, Reinhold-Hurek B (2003) Dechlorosoma suillum Achenbach et al. 2001 is a later
subjective synonym of Azospira oryzae Reinhold-Hurek and Hurek 2000. Int J Sys Evol
Microbiol 53:1139-1142
Wallace W, Beshear S, Williams D, Hospadar S, Owens M (1998) Perchlorate reduction by a
mixed culture in an up-flow anaerobic fixed bed reactor. J Indust Microbiol & Biotechnol
20:126-131
Wallace W, Ward T, Breen A, Attaway H (1996) Identification of an anaerobic bacterium which
reduces perchlorate and chlorate as Wolinella succinogenes. J Indust Microbiol 16:68-72
Wallner G, Amann R, Beisker W (1993) Optimizing fluorescent in situ hybridization with
rRNA-targeted oligonucleotide probes for flow cytometric identification of
microorganisms. Cytometry 14:136-143
56 Wolff J (1998) Perchlorate and the thyroid gland. Pharmacol Rev 50:89-106
Wu J, Unz RF, Logan BE (2001) Persistence of perchlorate and the relative numbers of
perchlorate- and chlorate-respiring microorganisms in natural waters, soils and
wastewater. Bioremed. J. 5:119-130
Xu J, Trimble JJ, Steinberg L, Logan BE (2004) Chlorate and nitrate reduction pathways are
separately induced in the perchlorate-respiring bacterium Dechlorosoma sp. KJ and the
chlorate-respiring bacterium Pseudomonas sp. PDA. Water Res 38:673-680.
Zhang H., Bruns MA, Logan BE (2002) Perchlorate reduction by a novel chemolithoautotrophic,
hydrogen-oxidizing bacterium. Environ Microbiol 4:570-576
57 80
70 0.34 L/m2-h 0.68 L/m2-h
60
50 g/L) µ
40
30 Perchlorate ( Perchlorate
20
10
0 0.1 0.3 0.5 0.7 0.9 1.1 1.3
Distance in reactor (m)
Figure 3.1 Perchlorate concentration profile in the reactor with two flow rates. Adapted from
Min et al. 2003, reprinted with permission.
58
70 10
9 60 Perchlorate
Acetate 8 Nitrate 50 Oxygen 7 (mg/L) 3 40 6 5 30 4
(ug/L), Acetate (mg/L) 20 3 4
2 DO (mg/L), NO ClO 10 1
0 0 0.1 0.3 0.5 0.7 0.9 1.1 1.3 Distance in Reactor (m)
Figure 3.2 Chemical profiles in the reactor of acetate, oxygen, nitrate, and perchlorate at 0.34 L
m-2s-1 flow rate. Adapted from Min et al. 2003, reprinted with permission
59
Figure 3.3 Inverted image of SYBR-green stained polyacrylamide gel showing ribosomal
intergenic spacer profiles of DNA extracted from the reactor biofilms.
60 A
B
C
Z = 0 µm Z = 15 µm Z = 30 µm
Z = 45 µm Z = 60 µm Z = 108 µm
Figure 3.4 FISH detection of Dechloromonas and Dechlorosoma species. Panel A and B: FITC- labelled Soma1035 hybridizing to Dechlorosoma sp. KJ and Cy3-labelled Monas1403 hybridizing to Dechloromonas sp. HZ, respectively. Panel C: Cy5-labelled EUB338, Soma1035, and Monas1403 hybridizing to cells in biofilms. Magenta cells were detected by both EUB338 and Monas1403. Cyan cells were detected by both EUB338 and Soma1035.
61 Portions of surface covered by bacteria (%)
0 102030405060 Depth from the surface of the biofilm ( 0
20
40
60
80
All bacteria 100 Dechloromonas
µ Dechlorosoma m) 120
Abundance of two perchlorate-reducing genera
as a percentage of all bacteria (%)
Depth from the surface of the biofilm ( 0 5 10 15 20 25 0
20
40
60
80
Dechloromonas 100 Dechlorosoma
µ m) 120
Figure 3.5 Relative abundance of bacteria hybridizing with EUB338, Monas, and Soma probes on the basis of total field area. Relative abundance of bacteria hybridizing with Monas and Soma probes on the basis of all bacteria. Data were obtained from COMSTAT program.
62 Chapter 4
H2 production by Clostridium acetobutylicum in a trickle-bed reactor
Abstract
H2 gas production through dark fermentation provides a renewable method of energy
production. Generated H2 can be captured and used in conventional fuel cells. H2 production in
continuous-flow stirred tank reactors, however, requires additional energy to mix reactor
contents in order to minimize H2 supersaturation in the liquid phase. Here we report the design and testing of an unsaturated flow (trickle-bed) H2-producing bioreactor. A column was packed with glass beads and inoculated with Clostridium acetobutylicum. Glucose in a minimal medium was fed at a flow rate of 1.6 mL/min (0.096 L/h) into the capped reactor, producing a hydraulic retention time of 2.1 min. Continuous H2 production ranged from 14.1 to 27.2 mL/h. The specific H2 production rate was 676 - 1265 mL/g-glucose per liter of reactor (total volume).
Gas-phase H2 concentrations of 70-79% were obtained at influent glucose concentrations ranging
from 1.0 to 10.5 g/L. At 3.3 g/L glucose concentration, biogas production and H2 concentration
were comparable between repeated tests. The major fermentation byproducts were acetate and
butyrate. The measured H2 yields indicated an overall conversion efficiency of 15-27% based on
a theoretical stoichiometry of 4 moles of H2 from 1 mole of glucose. Clogging of the column by
excreted biopolymers prevented the reactor to be operated longer than 60 hours. At one glucose
concentration, 3.3 g/L, cleaning the column enabled a stable operation for 72 hours and an
increased biogas production rate. The high H2 gas concentrations obtained in this study
demonstrated the feasibility of this unsaturated trickle-bed reactor for biological H2 production.
A likely application of this reactor is the generation of H2 gas from the pre-treatment of a high
carbohydrate-containing wastewater.
63 4.1 Introduction
H2 is considered to be a green energy carrier of the future. The only by-product of
reacting H2 with oxygen is water. No carbon dioxide or other green house gases are produced.
Most H2 is currently produced from nonrenewable sources such as oil, natural gas, and coal
(Benemann 1996; Van Ooteghem et al. 2002). H2 can also be produced from renewable sources
such as biomass. If H2 conversion efficiency could reach 60–80 % based upon a maximum
theoretical conversion of 12 mol-H2/mol-hexose, H2 production from wastewater could hold the
greatest potential for economical near-term H2 yield from renewable resources (Benemann
1996). However, no known fermentation pathways exist to achieve a conversion efficiency of
higher than 4 mol-H2/mol-hexose (Thauer et al. 1977). Although an enzyme assay was reported
to produce 11.6 mol-H2/mol-glucose by coupling enzymes from oxidative pentose phosphate
cycle and hydrogenase purified from Pyrococcus furiosus (Woodward et al. 2000), such a high conversion efficiency has not been reported in whole-organism studies.
Recent studies on biological H2 production has employed batch (Van Ginkel et al. 2001;
Logan et al. 2002; Oh et al. 2003), fed-batch (Chin et al. 2003), continuous-flow stirred tank
reactors (Fang and Liu 2002; Hussy et al. 2003), or saturated packed-bed column reactors
(Rachman et al. 1998). In batch tests, it has been found that constantly releasing biogas from the
system can increase H2 yields by as much as 40% (Logan et al. 2002). Typical conversion
efficiencies using continuous flow reactors are 1.9 – 2.4 mol-H2/mol-glucose (Lay 2001; Ueno et
al. 2001a; Fang and Liu 2002). In one study, continuous H2 production achieved a yield of 2.7
mol mol-H2/glucose using a mixed culture of Clostridium butyricum and Enterobacter aerogenes
(Yokoi et al. 2002). While stirring a reactor facilitates gas release from the liquid phase, it also
64 consumes considerable electric power. Mixing energy inputs in the range of 85 to 105 kW/1000 m3 are typically used for anaerobic waste treatment systems (Grady et al. 1999).
Trickle-bed reactors promote gas-liquid transfer through thin films obtained by flowing water over a support medium. It was postulated that H2 could diffuse out of biofilms and the
liquid film in the same way that oxygen diffuses into biofilms in a traditional trickling filter
(Logan 1993). Here we report the design and testing of such a trickle-bed column reactor for
biological H2 production. In this study we inoculated the reactor with a H2-producing gram
positive bacterium Clostridium acetobutylicum.
4.2 Materials and Methods
4.2.1 Medium and culture conditions
A stock culture of C. acetobutylicum ATCC 824 was stored at -80 oC. The working
culture was grown anaerobically in a synthetic medium that contained, in g L-1, glucose, 1;
NH4Cl, 0.2; KH2PO4, 1.8; K2HPO4, 2.40; MgSO4.7H2O, 0.1; FeCl3, 0.02; CaCl2, 0.01;
Na2BO7.H2O, 0.011; ZnCl2, 0.015; CoCl2.6H2O, 0.01; CuCl2.2H2O, 0.015; MnCl2.4H2O, 0.01.
The initial pH of the medium was 6.2. Cell density prior to inoculation was examined by optical
density measurements and by direct microscopic observation of acridine orange-stained cells
(Olympus BH-2). The inoculum was harvested in late-log phase of batch growth (O.D. ca. 0.2-
0.3). About 70 mL was used to inoculate the column reactor.
65 4.2.2 Reactor design and operation
A trickle-bed column reactor (Figure 1) was constructed based on modifications of a previous setup (Logan and LaPoint 2002) designed to minimize gas losses in the reactor effluent.
Water was pumped (Masterflex 7523-30, Cole Palmer Corp.) into the top of the reactor where it dripped onto the reactor packing. The main chamber of the reactor (25 cm long, 2.5 cm inside diameter, 0.123 L volume) was packed with autoclaved glass beads (3 mm diameter; 1200 m2/m3 calculated projected surface area) supported by a stainless steel mesh. The biogas produced in the reactor escaped through a gas port on the top of the reactor, with the flowrate of the gas monitored using a respirometer system (Challenge Environmental Systems AER-200 respirometer, Fayetteville, AR). The effluent exit was connected directly to a sealed Zip-Loc bag (1.5 L) containing 150 g brewer’s salt to halt biological activity in the collection system.
The reactor was operated in a constant temperature (30 oC) room. Oxygen was initially
removed from the column by purging with nitrogen gas for 30 s. The reactor was then operated
in batch mode until constant gas production was observed. At this time the column reactor was
switched to a continuous feed mode. Nitrogen-sparged sterile medium was fed at a flow rate of
1.6 mL/min into the top of the reactor. The pH was controlled using a phosphate buffer (see
medium composition). Headspace gas samples (0.25 mL) were periodically taken using a gas-
tight syringe. Effluent samples were taken at a point before the flow entered the collection bag.
After collection, aqueous (influent and effluent) samples were immediately centrifuged for 1 min at 8,000 ×g and the supernatant was stored at 4 oC for subsequent analysis. Steady state
operation was indicated by constant gas production and a stable headspace H2 concentration, and
verified by constant effluent glucose and volatile acids concentrations.
66 Four different influent glucose concentrations were chosen: 1.0, 3.3, 4.5, and 10.5 g/L.
When the influent glucose concentration was changed, the reactor was purged with nitrogen gas again in order to drop the H2 percentage to zero in the column headspace. These influent glucose
concentrations were comparable to those used in other H2-producing bioreactor studies (Mizuno
et al. 2000; Fang and Liu 2002; Iyer et al. 2004) and reflected the strength of sugar
concentrations in food processing industrial wastewaters (Van Ginkel and Logan in press). For
one condition, a glucose concentration of 3.3 g/L, the experiment was repeated twice, and a third
test involving column cleaning was included. To clean the column of excess biomass, fresh
medium was pumped from the column’s exit port to float the excess biomass. An ethanol-wiped
spatula was then used to mix the glass beads in the column and remove excess biofilm and other
biopolymers excreted by the microorganisms growing on the glass beads. The glucose
concentrations used in the test were in the following order: 4.5g/L, 3.3g/L(A),10g/L, 3.3g/L(C),
1g/L, 3.3g/L(B). At glucose concentrations of 4.5, 3.3(A), 10, and 3.3(C)g/L, fresh inoculum
was used, while at other glucose concentrations the culture on the glass beads were re-used.
4.2.3 Calculations
Volumetric H2 production rates RHV (as mL-H2/h) were calculated based upon total gas
production rates and steady state headspace H2 percentages. Molar H2 production rates RHM (as
RHV mmol-H2/h) were calculated using the ideal gas law as RHM = , where R = 0.0821 (L- RT
atm)/(mol-K), and T = 303 K. Conversion efficiency was calculated as (RHM/4RG)×100% where
RG (as mmol-glucose/h) is the glucose consumption rate (Table 4.1). The specific H2 production
67 1000R rate is S = HV , where V is the reactor’s total volume (0.123 L, 25 cm long, 2.5 cm inside 180RGV diameter).
The maximum theoretical production of H2 and CO2 were calculated using measured acetic and butyric acid concentrations assuming (Muller 2001) the following stoichiometry:
C6H12O6 + 2H2O 2CH3COOH + 4H2 + 2CO2 (1)
C6H12O6 CH3(CH2)2COOH + 2H2 + 2CO2 (2)
4.2.4 Determination of the reactor’s hydraulic retention time
The reactor’s hydraulic detention time was determined by spiking the feed line with a concentrated KCl solution (16 g/L), and measuring the conductivity of the reactor effluent (YSI
600XL; YSI Incorporated; Yellow Springs, OH, USA). The detention time was determined by using the front tracer curve in Figure 4.2 as previously described (Min et al. 2004).
4.2.5 Analytical procedures
H2 was measured with a gastight syringe (0.25 mL injection volume) and a gas chromatograph (SRI instruments, Torrence, CA) equipped with a thermal conductivity detector and a molecular sieve column (Alltech) with nitrogen gas as the carrier gas. Glucose was measured by using a phenol-sulfuric method for reducing sugars (Dubois et al. 1956). Acetate and butyrate were determined by gas chromatography (Agilent 6890N) with injector and flame ionization detector temperatures of 250 oC, with helium as a carrier gas at flow rates of 3.5 mL/min (60 oC) to 1.5 mL/min (240 oC). The oven temperature was programmed as follows: 60 oC for zero min, ramping to 120 oC at 20 oC/min, followed by ramping to 240 oC at 30 oC/min,
68 and hold at 240 oC for 3 min. The column used was DB-FFAP with dimensions of 30 m x 0.32 mm x 0.5 um.
4.3 Results and Discussion
At all glucose loading rates there was a high rate of biogas production after a lag phase ranging from 6-12 hours (when only the feeding glucose concentration changed but without re- inoculation) to 32-40 hours (when both the feed bottle and glass beads changed) (Figure 4.3).
The biogas production rate increased from 19±0 mL/h to 37±0 mL/h when the glucose loading rate was increased from 0.09 to 1.02 mmol/h (Table 4.1). The steady-state headspace H2 concentrations were 74±3%, 79±2%, 74±2%, and 70±1% for glucose concentrations of 10.5, 4.5,
3.3, and 1.0 g/L, respectively (Figure 4.4). The biogas production rates were comparable (28.2 vs. 27.4 mL/h) for the two experimental repeats at 3.3 g/L glucose concentration. Cleaning the column increased the gas production rate from 28 to 36 mL/h, resulting in an increase of H2 production from 20 to 26 mL/h. The effluent pH varied from 4.9 at 10 g/L to 5.6 at other glucose concentrations.
The conversion efficiencies ranged from 15 to 27 % based on a theoretical stoichiometry of 4 moles H2 from 1 mole of glucose (Table 4.1). Cleaning the column at 3.3 g/L glucose concentration increased the conversion efficiency from 15 to 19%. These conversion efficiencies were comparable to 24.2 % and 23 % reported in batch studies (Logan et al. 2002;
Oh et al. 2003), but less than 67.5 % (Yokoi et al. 2002) and 47.5 to 55% (Ueno et al. 2001b;
Fang and Liu 2002; Chin et al. 2003; Hussy et al. 2003) using fed-batch or continuous flow stirred reactors.
Table 4.2 lists the amount of acetate and butyrate production. Theoretical CO2 and H2 yields were calculated from volatile acid concentrations measured in the reactor effluent and
69 equations 1 and 2. The carbon recovery was 80 – 94 %, with the balance (6 – 20 %) assumed to have been converted to biomass. The predicted conversion efficiencies (60 – 72%) were much higher than the measured values (15 – 27%). This discrepancy could be due to several factors, including loss of H2 via homoacetogenesis (Oh et al. 2003), and loss of biogas through the collection bag. No methane was detected in the gas throughout the tests.
While the overall yield of H2 from glucose was low, the H2 production rate is comparable to that reported for several CSTR reactors. The H2 production rate, normalized by the reactor’s working volume, was found here to be 8.86 mmol-H2/(L-h). This is similar to 8.77 mL-H2/(L-h)
(Iyer et al. 2004) and 7.85 mL-H2/(L- h) (Mizuno et al. 2000), but higher than 2.52 mmol-H2/(L- h) (Fang and Liu 2002) (Table 4.3). This suggests that on the basis of reactor volume, the trickle-bed reactor was as effective as a CSTR.
Implications. The high H2 concentration obtained in this study demonstrated the feasibility of this novel unsaturated trickle-bed reactor for biological H2 production. A trickle- bed reactor has a low energy demand and is easy to scale-up due to simple construction needs.
For practical economical applications, organic waste materials need to be used as substrates for
H2 production to reduce cost. In addition, larger-scale reactors will be needed to increase the total removal rate of the influent organic matter. The performance data obtained from this study will be useful for anticipating potential H2 producing rates in larger systems.
Acknowledgements
This research was funded by NSF grant BES 01-24674 and by NSF (IGERT) grant DGE-
9972759, which supports the Penn State Biogeochemical Research Initiative for Education
(BRIE).
70 Table 4.1. Summary of H2 production rates and conversion efficiency (based upon a maximum production of 4 moles of H2 from every mole of glucose)
System conditions Influent glucose concentration (g/L)
10.5 4.5 3.3 1.0
Influent glucose (mmol/L) 58.4 25.0 18.3 5.6
Glucose loading rate (g/h) 1.02 0.43 0.32 0.09
Effluent glucose concentration (mmol/L) 45.7±1.7 9.8 ±1.2 4.2±0.5 0.1±0.0
Consumed glucose (mmol/L) 12.7±1.7 15.2±1.2 14.1±0.5 5.5±0.0
a Glucose consumption rate (RG) (mmol/L) 1.2±0.2 1.5±0.1 1.4±0.1 0.5±0.0
Biogas production rate (mL/h) 37±0 31±1 27±1 16±0
H2 (%) 74±4 79±2 74±2 70±1
Volumetric H2 production rate, RHV (mL/h) 27±1 25±1 20±1 11±1
Molar H2 production rate, RHM (mmol/h) 1.1±0.1 1.0±0.1 0.8±0.0 0.4±0.0
Conversion efficiency (%)b 22 17 15 15
Specific H2 production S: (mL-H2/L-g-glucose) 1007 761 676 1041 a based on a flowrate of 0.096 L/h b based on 4 mol-H2/mol-glucose
71 Table 4.2. Summary of volatile fatty acids production
System conditions Influent glucose concentration (g/L)
10.5 4.5 3.3 1.0
Consumed glucose, RG (mmol/h) 1.2±0.2 1.5±0.1 1.4±0.1 0.5±0.0
Consumed carbon (mmol/h) 7.3±1.0 8.8±0.7 8.1±0.3 3.0±0.1
Acetate (mmol/h) 1.2±0.2 1.5±0.1 1.3±0.1 0.4±0.1
Butyrate (mmol/h) 0.5±0.1 0.5±0.1 0.6±0.0 0.2±0.1
a. Predicted CO2 (mmol/h) 2.3±0.4 2.5±0.3 2.5±0.1 0.8±0.2
Carbon recovered in acetate, butyrate, and CO2 6.9±1.1 7.6±1.0 7.5±0.4 2.4±0.3 (mmol/h) Carbon recovery (%) 94 87 92 80
b. Predicted H2 (mmol/h) 3.5±0.5 4.1±0.4 3.8±0.2 1.2
Predicted conversion efficiency c. (%) 72 69 70 60
a. Based on equation 1 and 2, assuming the production of 1 mole of CO2 with the production of 1 mole of acetate and 2 moles of CO2 with the production of 1 mole of butyrate b. Based on equation 1 and 2, assuming the production of 2 moles of H2 with the production of 1 mole of acetate or butyrate. c. Predicted conversion efficiency calculated as (predicted H2)/(4RG) x 100%
72 Table 4.3. Comparison of H2 production rates in the trickle-bed reactor with those reported using a CSTR
Reactor configuration Comparison Items Trickle-bed CSTR CSTR CSTR reactor (Mizuno et (Fang and (Iyer et al. Reference This study al. 2000) Liu 2002) 2004) Reactor detention time 2.1 min 8.5 h 6 h 10 h Volumetric H production rate 2 27.2 457 192 436 (mL/h)
Molar H2 production rate (mmol/h) 1.09 18.06 7.57 17.53
Reactor working volume (L) 0.123 2.3 3 2
Normalized H production rate 2 8.86 7.85 2.52 8.77 (mmol-H2/L-h)
73
Medium in Gas out to a bubble meter
Syringe sampling port
Glass beads for biofilm buildup
Stainless steel mesh
Zip-loc bag for collecting waste
Figure 4.1. A picture and a schematic drawing of the trickle-bed reactor
74 4500
4000 )
3500
3000 2500
2000
Conductivity (uS/cm 1500
1000
0 60 120 180 240 300
Time (sec)
Figure 4.2 Determination of reactor detention time by tracer study. Solid symbols are measured conductivity; open circles are symmetrical with the front half of the actual curve.
75 900 10g/L 800 4.5g/L 700 3.3g/L(A) 3.3g/L(B) 600 3.3g/L(C) 500 1.0g/L
400 Biogas (mL) Biogas 300
200
100
0 0 1224364860 Time (h)
Figure 4.3 Biogas productions over time. Solid symbols were used for regression. The slopes of the regression lines represented gas production rates and were summarized in Table 4.1. The operation with column cleaning was indicated as 3.3g/L(C).
90% 10g/L 80% 4.5g/L 3.3g/L(A) 70% 3.3g/L(B) 60% 1.0g/L 3.3g/L(C) 50% (%) 2
H 40%
30%
20%
10%
0% 0 1224364860 Time (h)
Figure 4.4 Headspace H2 concentrations over time
76 3600 1000
900 3000 800 700 2400 600 1800 Glu in 500 Glu out 400
Glucose (mg/L) Glucose 1200 Acetate Butyrate 300 200 (mg/L) Butyrate or Acetate 600 100 0 0 0 4 8 12162024283236 Time (h)
Figure 4.5 Glucose, acetate, and butyrate concentration profiles during the column operation at the 3.3 g/L (B) influent glucose concentration.
77 4.4 References
Benemann J (1996) H2 biotechnology: Progress and prospects. Nat Biotechnol 14:1101-1103
Chin HL, Chen ZS, Chou CP (2003) Fedbatch operation using Clostridium acetobutylicum
suspension culture as biocatalyst for enhancing H2 production. Biotechnol Prog 19:383-
388
Dubois M, Gilles K, Hamilton J, Rebers P, Smith F (1956) Colorimetric method for
determination of sugars and related substances. Anal Chem 28:351-356
Fang HH, Liu H (2002) Effect of ph on H2 production from glucose by a mixed culture.
Bioresour Technol 82:87-93
Grady CPL, Daigger GT, Lim HC (1999) Biological wastewater treatment, 2nd rev. and expand
edn. Marcel Dekker, New York
Hussy I, Hawkes FR, Dinsdale R, Hawkes DL (2003) Continuous fermentative H2 production
from a wheat starch co-product by mixed microflora. Biotechnol Bioeng 84:619-626
Iyer P, Bruns MA, Zhang H, van Ginkel S, Logan BE (2004) H2-producing bacterial
communities from a heat-treated soil inoculum. Appl Microbiol Biotechnol Online
First:DOI: 10.1007/s00253-00004-01666-00257
Lay JJ (2001) Biohydrogen generation by mesophilic anaerobic fermentation of microcrystalline
cellulose. Biotechnol Bioeng 74:280-287
Logan BE (1993) Oxygen-transfer in trickling filters. J Environ Eng-ASCE 119:1059-1076
Logan BE, LaPoint D (2002) Treatment of perchlorate- and nitrate-contaminated groundwater in
an autotrophic, gas phase, packed-bed bioreactor. Water Res 36:3647-3653
Logan BE, Oh S, Kim IS, Van Ginkel S (2002) Biological H2 production measured in batch
anaerobic respirometers. Environ Sci Technol 36:2530-2535
78 Min B, Evans PJ, Chu AK, Logan BE (2004) Perchlorate removal in sand and plastic media
bioreactors. Water Res 38:47-60
Mizuno O, Dinsdale R, Hawkes FR, Hawkes DL, Noike T (2000) Enhancement of H2 production
from glucose by nitrogen gas sparging. Bioresource Technol 73:59-65
Muller V (2001) Bacterial fermentation. Encylopedia of Life Sciences
Oh SE, Van Ginkel S, Logan BE (2003) The relative effectiveness of ph control and heat
treatment for enhancing biohydrogen gas production. Environ Sci Technol 37:5186-5190
Rachman MA, Nakashimada Y, Kakizono T, Nishio N (1998) H2 production with high yield and
high evolution rate by self-flocculated cells of Enterobacter aerogenes in a packed-bed
reactor. Appl Microbiol Biot 49:450-454
Thauer, RK., Jungermann K, Decker K. (1977). Energy conservation in chemotrophic anaerobic
bacteria. Bacteriol Rev 41:100-80.
Ueno Y, Haruta S, Ishii M, Igarashi Y (2001a) Characterization of a microorganism isolated
from the effluent of H2 fermentation by microflora. J Biosci Bioeng 92:397-400
Ueno Y, Haruta S, Ishii M, Igarashi Y (2001b) Microbial community in anaerobic H2-producing
microflora enriched from sludge compost. Appl Microbiol Biotechnol 57:555-562
Van Ginkel S, Sung SW, Lay JJ (2001) Biohydrogen production as a function of pH and
substrate concentration. Environ Sci Technol 35:4726-4730
Van Ooteghem SA, Beer SK, Yue PC (2002) H2 production by the thermophilic bacterium
Thermotoga neapolitana. Appl Biochem Biotechnol 98-100:177-189
Woodward J, Orr M, Cordray K, Greenbaum E (2000) Biotechnology - enzymatic production of
biohydrogen. Nature 405:1014-1015
79 Yokoi H, Maki R, Hirose J, Hayashi S (2002) Microbial production of H2 from starch-
manufacturing wastes. Biomass Bioenergy 22:389-395
80 Chapter 5
Microbial community response to hydraulic retention time (HRT) and substrate concentration in a chemostat H2-producing reactor
5.1 Introduction
Hydrogen gas is currently produced from nonrenewable sources such as natural gas, oil,
and coal (Benemann 1996; Van Ooteghem et al. 2002). Hydrogen can also be produced through electrolysis of water, but the cost of this simple process is higher than those based on fossil fuels
(Oh et al. 2003; Logan 2004). More sustainable methods are needed to produce hydrogen. One such method is to produce hydrogen with microorganisms by fermentation of organic matter from waste materials.
Pure cultures of microorganisms such as Enterobacter aerogenes, Clostridium butyricum, and Clostridium acetobutylicum have been used to produce hydrogen (Karube et al. 1982;
Heyndrickx et al. 1987; Rachman et al. 1997; Rachman et al. 1998; Chin et al. 2003). Studies using these microorganisms were performed under aseptic conditions with reported conversion efficiencies between 1.1 to 2.4 mol-H2/mol-glucose, based upon a maximum conversion of 4
moles of hydrogen per mole of glucose.
To produce hydrogen from non-sterile substrates such as industrial and municipal
wastewater, the use of pure culture conditions will likely be impossible. Biological hydrogen
production from mixed consortia have been reported in batch or continuous flow reactors (Lay
2001; Ueno et al. 2001; Van Ginkel et al. 2001; Fang and Liu 2002; Logan et al. 2002; Oh et al.
81 2003). Conversion efficiencies reported from these studies have varied from 0.7 to 2.7 mol-
H2/mol-glucose. Although significant yields of H2 have been obtained using mixed cultures in
batch and continuous systems, very little information is available on the microbial species that
exist in these reactors. In a mixed culture fermentation system, hydrogen-consuming organisms
could co-exist with hydrogen producing microorganisms. These hydrogen-consuming microbes
include nitrate-reducing bacteria, sulfate-reducing bacteria, CO2-reducing homoacetogenic
bacteria, and methanogens. Some of these hydrogen consumers can be eliminated using a heat-
shock process (Lay 2000; Van Ginkel et al. 2001; Logan et al. 2002), selecting for spore-forming
microbes. Using treated sludge as inocula, it has been found that Clostridium,
Sporolactobacillus, Streptococcus, and Streptococcus spp. were present in hydrogen-producing
microbial communities (Ueno et al. 2001; Fang et al. 2002). In the current study our objective
was to identify the bacterial populations that became established in a continuous flow bioreactor
operated at four different hydraulic retention times (HRT) (1, 2.5, 5, and 10h) and glucose
concentrations (2.5, 5, 7.5, and 10 g/L). The choice of this experimental matrix was based on the
COD range of sugar-containing industrial wastewaters, as well as for the purpose of testing the
effect of H2 production rate on H2 yields from glucose.
5.2 Materials and Methods
Reactor inoculation and operation. Surface soil from tomato plots on the Penn State
University Park campus was sampled, sieved, and dried for 2 h at 105°C to kill vegetative
bacteria and stored at -20oC as described previously (Logan et al. 2002). Inocula consisted of a
10-g samples added to 2 L of medium. A continuous flow stirred tank reactor (BioFlo 110, New
Brunswick Sciences) with a working volume of 2 L was inoculated and operated at 30 oC in
82 batch mode until gas production started and the redox potential dropped below 200, after which the reactor was operated in continuous mode. In order to suppress methanogenic activity, pH was maintained at 5.5 with 1 M KOH. Establishment of steady-state conditions was assessed by measuring total gas and H2 production and determining that production rates were constant over a period of at least three HRTs. Synthetic waste water feed consisted of the following
-1 components in g L concentrations: glucose (variable); NH4Cl (0.88); KH2PO4 (0.40); K2HPO4
(0.40); MgSO4.7H2O (0.48); FeCl3 (0.075); NiSO4 (0.048); CaCl2 (0.075); Na2BO7.H2O (0.011);
ZnCl2 (0.035); CoCl2.6H2O (0.032); CuCl2.2H2O (0.015); MnCl2.4H2O (0.045).
Chemical assays. The biogas production was continuously monitored with a bubble meter
calibrated according to the manufacturer’s instructions (Challenge Environmental Systems AER-
200 respirometer, Fayetteville, AR). Glucose was measured by using a published phenol-sulfuric
acid method (Dubois 1956). Headspace hydrogen was measured with a gas chromatograph
equipped with a thermal conductivity detector and a molecular sieve column (Molesieve 5A
80/100 6’×1/8’×0.085, Alltech) with nitrogen as the carrier gas.
DNA extraction. Of all 16 conditions (4 HRTs and 4 glucose concentrations), extensive
flocculation was observed under 13 conditions. Only flocculated samples were used to extract
DNA since as an initial attempt to unravel the bioreactor microbial assemblages, flocculated
samples suggested high biomass concentration thus high H2 production rates. Cells were lysed
with a 2-min beadbeater treatment (BioSpec, Bartlesville, OK) and DNA was purified with an
UltraClean Soil DNA Kit according to the manufacturer’s protocol (MoBio Laboratories,
Carlsbad, CA). DNA yields were between 500 ng and 2 µg per 2 mL sample. DNA was
quantified and its purity verified spectrophotometrically with a Lambda 40 UV/VIS
spectrophotometer (Perkin Elmer, Norwalk, CT).
83 PCR and RISA fingerprinting. The procedure was adapted from a previously published paper (Zhang et al. 2002). Ribosomal intergenic spacer analysis was referred to as RISA fingerprinting. DNA was amplified by polymerase chain reaction (PCR) with the bacteria- specific 16S rDNA primer set 926f and 115r (Lane 1991). PCR buffer (Promega, Madison,
WI), 2.5 mM MgCl2, 1.25 U of Taq (Promega), 30 pmol of each primer and 0.4 mM of each
deoxynucleoside triphosphate (dNTP) and 10 ng of template DNA were used per reaction in a
final volume of 50 µl. Amplification was performed according to the following conditions in a
thermocycler (GeneAmp PCR 2400, Perkin Elmer, Norwalk, CT): 5 min at 94°C; followed by 30
cycles of 30 sec at 94°C, 30 sec at 56°C and 1 min at 72°C; followed by a final extension at 72°C
for 7 min.
PCR-amplified DNA products were separated by electrophoresis in 1.5 % agarose gels.
The gel was prepared and run in 1X TAE buffer (pH 8.0) for 2 hours at 90 Volts. The gel was
stained with ethidium bromide for 30 minutes and rinsed with deionized water for 20 min. The
gel image was acquired with an Epi Chemi II Darkroom unit (UVP Laboratory Products, Upland,
CA). DNA bands were excised and eluted from the gel by the crush and soak procedure
(Sambrook et al. 1989). Eluted DNA was ligated into a plasmid vector (TOPO TA, Invitrogen)
for cloning into competent E. coli cells. DNA sequences were obtained from at least two purified
transformants for each band (ABI Hitachi 3100). Sequence editing and alignment were
conducted by using EditSeq and SeqMan programs (Lasergene). The CHIMERA_CHECK
(Cole et al. 2003) and Bellerophon (Hugenholtz and Huber 2003) programs were used to
evaluate sequences for chimeras. Closest relatives for 16S rRNA gene sequences were identified
by database searches with BLAST (Altschul et al. 1990).
84 5.3 Results and Discussion
Overall the microbial community identified in the reactor was relatively simple with not more than five distinct bands per lane (Figure 5.1). Similar banding patterns were observed for all HRT and glucose concentration conditions except for the 2.5 g/L glucose concentration. A heavily stained band appeared in all 10, 7.5, and 5 g/L glucose conditions, suggesting the presence of a common microorganism. Sequencing results showed this band was a close relative
(over 97%) of Clostridium acidisoli (Table 5.1). On the other hand, at the 2.5 g/L glucose concentration, a different banding pattern was observed. The community seemed more diverse than those obtained at other glucose concentrations. Sequencing results revealed that instead of
Clostridium acidisoli, Selenomonadaceae spp. were present in the reactor.
The detection of sequences belonging to Clostridium spp. at all but 2.5 g/L glucose concentrations was not surprising, since spore-forming bacteria were likely to have survived the soil heat treatment. In the current study I only chose to sequence the most heavily stained bands which were identified as close relatives of C. acidisoli, which belongs to Clostridium Cluster 1
(Collins et al. 1994). The temperature optima reported in the literature for C. acidisoli (25°-
30°C), an acid-tolerant isolate from forest litter (Kuhner et al. 2000), matched well with the reactor operating temperature (30°C). Our results were also in agreement to a previous study that found only Clostridiaceae was in a H2-producing reactor at 10 h HRT (Iyer et al. 2004).
Sequences closely related to Clostridium spp. were also found to account for 64.6% of the total
clones in a H2-producing bioreactor using the 16S rDNA clone library approach (Fang et al.
2002). The only sequence outside of Clostridiaceae cluster I retrieved in the current study,
Selenomonadaceae sp. S90, was isolated from anoxic soil (Chin et al. 1999). A previously
published review listed Enterobactreiaceae spp., Lactobacillacaeae spp., and Bacillaceae spp. as
85 the most common H2-producing facultative anaerobes (Nandi and Sengupta 1998). In the current study, no microorganisms from the above classes were found in the reactor.
Previous community profiling studies have used 16S rDNA clone libraries (Fang et al.
2002) and denaturing gradient gel electrophoresis (DGGE) (Ueno et al. 2001). In contrast to clone libraries, DNA fingerprinting techniques such as RISA and DGGE offer more convenient and comprehensive means for tracking of microbial populations in bioreactors, therefore suitable for use in monitoring community shifting. The RISA fingerprinting method used in this study offers more 16S rDNA sequence information than DGGE because nearly 600 base pairs were sequenced, instead of ca. 200 base pairs usually obtained by using DGGE.
86 Table 5.1 Phylogenetic summary of hydrogen-producing community from cloning and sequencing results (only 16S rDNA portion up to “GATCAC” were used in BLAST search).
Reactor condition ( glucose GenBank closest relative Similarity (%) Accession concentration and HRT) 5g/L-10h Clostridium acidisoli strain CK74 99 M23930 5g/L-1h Clostridium acidisoli strain CK74 97 M23930 5g/L-2.5h Clostridium acidisoli strain CK74 98 AJ237756 5g/L-5h Clostridium acidisoli strain CK74 99 AJ237756 2.5g/L-10h Selenomonadaceae strain SB90 97 AJ229242
87
Figure 5.1 Inverted image of ethidium bromide-stained agarose gels showing DNA bands in
RISA profiles. Lane 1, DNA marker. Lanes 2 to 14, samples from bioreactors operated at four different HRTs (1, 2.5, 5, and 10 h) and four different glucose feed concentrations (2.5, 5, 7.5, and 10 g/L).
88 5.4 References
Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search
tool. J Mol Biol 215:403-410
Benemann J (1996) Hydrogen biotechnology: Progress and prospects. Nat Biotechnol 14:1101-
1103
Chin HL, Chen ZS, Chou CP (2003) Fedbatch operation using Clostridium acetobutylicum
suspension culture as biocatalyst for enhancing hydrogen production. Biotechnol Prog
19:383-388
Chin K-J, Hahn D, Hengstmann U, Liesack W, Janssen PH (1999) Characterization and
identification of numerically abundant culturable bacteria from the anoxic bulk soil of
rice paddy microcosms. Appl Environ Microbiol 65:5042-5049
Cole JR et al. (2003) The ribosomal database project (RDP-II): Previewing a new autoaligner
that allows regular updates and the new prokaryotic taxonomy. Nucl Acids Res 31:442-
443
Collins MD et al. (1994) The phylogeny of the genus Clostridium: Proposal of five new genera
and eleven new species combinations. Int J Syst Bacteriol 44:812-826
Fang HH, Liu H (2002) Effect of ph on hydrogen production from glucose by a mixed culture.
Bioresour Technol 82:87-93
Fang HH, Zhang T, Liu H (2002) Microbial diversity of a mesophilic hydrogen-producing
sludge. Appl Microbiol Biotechnol 58:112-118
89 Heyndrickx M, Devos P, Thibau B, Stevens P, Deley J (1987) Effect of various external factors
on the fermentative production of hydrogen gas from glucose by clostridium-butyricum
strains in batch culture. Syst Appl Microbiol 9:163-168
Hugenholtz P, Huber T (2003) Chimeric 16s rDNA sequences of diverse origin are accumulating
in the public databases. Int J Syst Evol Microbiol 53:289-293
Iyer P, Bruns MA, Zhang H, van Ginkel S, Logan BE (2004) H2-producing bacterial
communities from a heat-treated soil inoculum. Appl Microbiol Biotechnol Online
First:DOI: 10.1007/s00253-00004-01666-00257
Karube I, Urano N, Matsunaga T, Suzuki S (1982) Hydrogen production from glucose by
immobilized growing cells of Clostridium butyricum. Appl Microbiol Biotechnol 16:5-9
Kuhner C, Matthies C, Acker G, Schmittroth M, Gossner A, Drake H (2000) Clostridium akagii
sp. nov. and Clostridium acidisoli sp. nov.: acid-tolerant, N2-fixing clostridia isolated
from acidic forest soil and litter. Int J Syst Evol Microbiol 50:873-881
Lane DJ (1991) 16s/23s rrna sequencing. In: Stackebrandt E, Goodfellow M (eds) Modern
microbiological methods. Wiley, Chichester ; New York
Lay JJ (2000) Modeling and optimization of anaerobic digested sludge converting starch to
hydrogen. Biotechnol Bioeng 68:269-278
Lay JJ (2001) Biohydrogen generation by mesophilic anaerobic fermentation of microcrystalline
cellulose. Biotechnol Bioeng 74:280-287
Logan BE (2004) Feature article: Biologically extracting energy from wastewater: Biohydrogen
production and microbial fuel cells. Environ Sci Tech 38:160A-167A
Logan BE, Oh S, Kim IS, Van Ginkel S (2002) Biological hydrogen production measured in
batch anaerobic respirometers. Environ Sci Technol 36:2530-2535
90 Nandi R, Sengupta S (1998) Microbial production of hydrogen: An overview. Crit Rev
Microbiol 24:61-84
Oh SE, Van Ginkel S, Logan BE (2003) The relative effectiveness of pH control and heat
treatment for enhancing biohydrogen gas production. Environ Sci Technol 37:5186-5190
Rachman MA, Furutani Y, Nakashimada Y, Kakizono T, Nishio N (1997) Enhanced hydrogen
production in altered mixed acid fermentation of glucose by enterobacter aerogenes. J
Ferment Bioeng 83:358-363
Rachman MA, Nakashimada Y, Kakizono T, Nishio N (1998) Hydrogen production with high
yield and high evolution rate by self-flocculated cells of Enterobacter aerogenes in a
packed-bed reactor. Appl Microbiol Biot 49:450-454
Sambrook J, Fritsch EF, Maniatis T (1989) Molecular cloning : A laboratory manual, 2nd edn.
Cold Spring Harbor Laboratory Press, Plainview, N.Y.
Ueno Y, Haruta S, Ishii M, Igarashi Y (2001) Microbial community in anaerobic hydrogen-
producing microflora enriched from sludge compost. Appl Microbiol Biotechnol 57:555-
562
Van Ginkel S, Sung SW, Lay JJ (2001) Biohydrogen production as a function of pH and
substrate concentration. Environ Sci Technol 35:4726-4730
Van Ooteghem SA, Beer SK, Yue PC (2002) Hydrogen production by the thermophilic
bacterium Thermotoga neapolitana. Appl Biochem Biotechnol 98-100:177-189
Zhang H, Bruns MA, Logan BE (2002) Perchlorate reduction by a novel chemolithoautotrophic,
hydrogen-oxidizing bacterium. Environ Microbiol 4:570-57
91 APPENDIX Data used to generate figures in chapter 2 to chapter 4
Data for Figure 2.1 Perc(mM)Time (h) 5.1500 0.0000 5.0460 10.0000 2.9700 24.0000 1.9800 28.0000 5.1500 28.0000 3.1200 32.0000 0.0000 40.0000 5.1500 40.0000 3.2510 41.0000 2.6850 42.0000 0.6820 43.0000 0.0000 46.0000 5.1500 46.0000 0.0000 48.0000
Data for Figure 2.3 Time(h) OD.h OD.n Clh Cln SDCln SDClh 0.0000 0.0310 0.0330 0.2900 0.1512 2.0000e-3 0.2700 12.0000 0.0420 0.0350 0.9700 0.1581 2.0000e-3 0.2900 17.5000 0.0640 0.0310 1.2700 0.1611 1.0000e-3 0.2600 25.0000 0.1040 0.0330 2.6800 0.1806 4.0000e-3 0.2100 32.5000 0.2060 0.0340 6.6500 0.1826 6.0000e-3 0.2400 35.5000 0.2680 0.0370 7.6500 0.1977 2.0000e-3 0.2900 38.5000 0.3070 0.0330 8.6000 0.2191 2.0000e-3 0.5800 45.5000 0.3090 0.0300 8.8300 0.2208 5.0000e-3 0.4300 58.0000 0.3090 0.0280 9.7200 0.2219 3.0000e-3 0.6100
92 70.0000 0.3060 0.0280 10.1900 0.2267 1.0000e-3 0.4200
Data for Figure 3.5 1.0000 111.0000 20.8035 0.3216 0.8976 1.5458 4.3146 2.0000 108.0000 21.4039 0.2579 0.6638 1.2048 3.1011 3.0000 105.0000 21.6984 0.3967 0.4871 1.8284 2.2450 4.0000 102.0000 21.8700 0.6577 0.3273 3.0071 1.4966 5.0000 99.0000 21.7972 0.7786 0.2422 3.5719 1.1113 6.0000 96.0000 21.6637 0.8076 0.1808 3.7278 0.8347 7.0000 93.0000 21.1731 0.7992 0.1266 3.7745 0.5982 8.0000 90.0000 20.8549 0.7381 0.0927 3.5394 0.4445 9.0000 87.0000 20.6211 0.6699 0.0912 3.2484 0.4421 10.0000 84.0000 20.8324 0.5531 0.0725 2.6551 0.3479 11.0000 81.0000 21.2685 0.4448 0.0599 2.0913 0.2816 12.0000 78.0000 21.4645 0.3452 0.0336 1.6084 0.1564 13.0000 75.0000 21.9093 0.2876 0.0172 1.3128 0.0784 14.0000 72.0000 22.5479 0.2350 0.0114 1.0422 0.0508 15.0000 69.0000 23.2334 0.1953 6.8665e-3 0.8407 0.0296 16.0000 66.0000 23.9868 0.1553 3.8147e-3 0.6473 0.0159 17.0000 63.0000 24.4049 0.1377 4.1962e-3 0.5643 0.0172 18.0000 60.0000 25.2220 0.1228 0.0103 0.4870 0.0408 19.0000 57.0000 25.9060 0.1240 0.0153 0.4786 0.0589 20.0000 54.0000 26.7132 0.1431 8.7738e-3 0.5355 0.0328 21.0000 51.0000 27.7378 0.1190 7.6294e-3 0.4291 0.0275 22.0000 48.0000 28.9295 0.1163 9.5367e-3 0.4022 0.0330 23.0000 45.0000 30.2433 0.1137 0.0114 0.3759 0.0378 24.0000 42.0000 31.4522 0.1064 0.0107 0.3384 0.0340 25.0000 39.0000 32.6817 0.1057 0.0278 0.3233 0.0852 26.0000 36.0000 33.8806 0.3613 0.0137 1.0663 0.0405 27.0000 33.0000 34.7134 0.4498 0.0134 1.2956 0.0385 28.0000 30.0000 35.4164 0.6794 9.9182e-3 1.9183 0.0280
93 29.0000 27.0000 36.2877 0.9361 7.2479e-3 2.5797 0.0200 30.0000 24.0000 37.3894 1.1040 6.8665e-3 2.9526 0.0184 31.0000 21.0000 38.4094 1.4885 0.0126 3.8753 0.0328 32.0000 18.0000 39.5592 1.6487 0.0118 4.1677 0.0299 33.0000 15.0000 40.7459 1.8795 0.0156 4.6127 0.0384 34.0000 12.0000 42.6281 2.1179 0.0118 4.9684 0.0277 35.0000 9.0000 46.1056 3.1513 0.1194 6.8350 0.2590 36.0000 6.0000 47.8401 4.6444 0.1362 9.7082 0.2847 37.0000 3.0000 49.1615 7.6241 0.1904 15.508 0.3872 38.0000 0.0000 50.7797 11.618 0.1762 22.880 0.3471
94 Data for Figure 4.2 41 192 3911 fact # t (sec) C (uS/cm) 42 196.8 3306 1 0 1152 43 201.6 3394 2 4.8 1543 44 206.4 3005 3 9.6 1995 45 211.2 3110 4 14.4 2306 47 220.8 3172 5 19.2 2247 48 225.6 2814 7 28.8 2751 50 235.2 2782 8 33.6 2864 51 240 2623 11 48 3069 52 244.8 2559 13 57.6 3324 53 249.6 2352 16 72 3508 55 259.2 2296 17 76.8 3621 56 264 2301 20 91.2 3883 58 273.6 2280 21 96 3834 60 283.2 2286 22 100.8 3830 23 105.6 4105
25 115.2 4154
27 124.8 4181 28 129.6 4031 29 134.4 4095
32 148.8 3906
34 158.4 4032 35 163.2 4072 36 168 3642
38 177.6 3750
95
Data for Figure 4.3
Elapse T 10g/L dgas/dt time(h) 4.5g/L time (h) 3.3g/L Hours 1.04g/L 3.3g rpt 0 0 0 0 0 9.18 0 0 0.927111435 1 0 0 0.3 0.4 1.333333333 0.5 9.18 0.5 0.927111435 2 0 0 0.5 0.8 2 1 9.18 1 0 4.372351807 3 0 0 1 1.47 1.34 1.5 9.18 1.5 0 4.372351807 4 0 0 1.5 2.13 1.32 2 9.18 2 0.07628109 4.372351807 5 0 0 2 3.06 1.86 2.5 9.18 2.5 0.07628109 4.372351807 6 0 0 2.5 4.26 2.4 3 9.18 3 0.781252877 4.372351807 7 0 0 3 5.59 2.66 3.5 9.18 3.5 0.781252877 4.372351807 8 0 0 3.5 7.05 2.92 4 9.18 4 0.781252877 4.372351807 9 0 0 4 8.78 3.46 4.5 9.18 4.5 0.781252877 7.866906441 10 0 0 4.5 9.98 2.4 5 9.18 5 0.781252877 15.42255925 11 0 0 5 9.98 0 5.5 9.18 5.5 0.781252877 28.22036849 12 0 0 5.5 11.44 2.92 6 9.18 6 0.781252877 45.47098036 13 12.4215 12.4215 6 13.7 4.52 6.5 9.18 6.5 0.781252877 59.48364063 14 35.456 23.0345 6.5 15.96 4.52 7 9.18 7 0.927111435 83.40353899 15 99.688 64.232 7 18.09 4.26 7.5 9.18 7.5 0.927111435 105.9584967 16 132.932 33.244 7.5 19.82 3.46 8 9.18 8 4.372351807 110.2313203 17 166.92 33.988 8 21.28 2.92 8.5 9.18 8.5 4.372351807 116.0438817 18 204.652 37.732 8.5 21.28 0 9 9.18 9 4.372351807 120.4951397 19 226.9 22.248 9 21.28 0 9.5 9.18 9.5 4.372351807 139.9777982 20 257.348 30.448 9.5 21.28 0 10 9.58 10 4.372351807 150.9658091 21 303.212 45.864 10 21.28 0 10.5 11.04 10.5 4.372351807 167.225412 22 351.084 47.872 10.5 21.28 0 11 14.1 11 4.372351807 182.9614729 23 394.74 43.656 11 21.28 0 11.5 16.36 11.5 7.866906441 207.7044403 24 424.44 29.7 11.5 21.28 0 12 19.68 12 7.866906441 227.3603215 25 462.28 37.84 12 21.42 0.28 12.5 32.59 12.5 30.20646351 238.6629124 26 503.69 41.41 12.5 23.01 3.18 13 32.85 13 47.52307751 251.3241662 27 538.23 34.54 13 23.01 0 13.5 32.85 13.5 67.96468135 255.6111109 28 578.31 40.08 13.5 23.01 0 14 32.85 14 83.13438774 267.7491054 29 610.52 32.21 14 23.14 0.26 14.5 32.85 14.5 107.1200844 285.2128435 30 647.92 37.4 14.5 23.14 0 15 32.85 15 128.1841676 306.2525062 31 682.28 34.36 15 23.14 0 15.5 32.85 15.5 132.0417598 319.4305557 32 717.84 35.56 15.5 23.14 0 16 32.85 16 137.2300534 330.0300028
96 33 751.58 33.74 16 23.14 0 16.5 32.85 16.5 141.1593643 340.1370131 34 785.16 33.58 16.5 23.14 0 16.78 32.85 17 157.9526495 354.6794702 35 841.34 56.18 17 23.14 0 17 32.85 17.5 167.1635155 366.9996108 36 857.62 16.28 17.5 23.14 0 17.5 32.85 18 180.4925384 387.0272685 37 892.47 34.85 18 23.14 0 18 32.85 18.5 193.0866655 406.8881782 38 954.64 62.17 18.5 23.28 0.28 18.5 32.85 19 212.3571473 423.6564078 39 977.96 23.32 19 24.47 2.38 18.5 32.85 19.5 227.2576716 434.0700156 40 1000.67 22.71 19.5 25.54 2.14 19 32.85 20 235.6791964 443.568619 20 26.87 2.66 19.5 32.85 20.5 244.9955717 457.02506 20.5 28.46 3.18 20 32.85 21 248.1231823 468.3207958 21 29.93 2.94 20.5 32.85 21.5 256.908556 488.1324232 21.5 32.32 4.78 21 32.85 22 269.3762828 507.2694271 22 34.72 4.8 21.5 32.85 22.5 284.1464832 510.9350865 22.5 37.38 5.32 22 32.85 23 293.2681432 536.7480982 23 41.23 7.7 22.5 32.85 23.5 300.5367697 546.3963798 23.5 44.29 6.12 23 32.85 24 307.4135223 560.7690286 24 47.75 6.92 23.5 32.85 24.5 317.2192873 574.7763453 24.5 50.94 6.38 24 32.85 25 325.4481068 596.9710593 25 55.6 9.32 24.5 32.85 25.5 338.6793522 614.7327911 25.5 55.6 0 25 32.85 26 351.6323724 624.9928487 26 55.6 0 25.5 32.85 26.5 362.4456413 636.5234896 26.5 55.6 0 26 32.85 27 369.1070896 642.8994131 27 55.6 0 26.5 32.85 27.5 375.1484505 651.5362722 27.5 55.6 0 27 32.85 28 383.6520804 667.5609202 28 55.6 0 27.5 32.85 28.5 390.7420306 686.9448671 28.5 55.6 0 28 32.85 29 403.0749313 699.1259323 29 55.6 0 28.5 32.85 29.5 414.8695526 708.9444055 29.5 55.6 0 29 32.85 30 417.1159934 718.3232659 30 56.53 1.86 29.5 32.85 30.5 432.8231013 731.8449832 30.5 58.12 3.18 30 32.85 31 438.6452256 743.3240244 31 58.12 0 30.5 32.85 31.5 447.2708156 762.0278924 31.5 58.12 0 31 32.85 32 455.6241251 780.6257811 32 58.26 0.28 31.5 32.85 32.5 468.7567179 796.3636038 32.5 58.39 0.26 32 32.85 33 479.1785088 33 58.39 0 32.5 33.78 33.5 485.1642959 33.5 59.32 1.86 33 34.05 34 491.8621151 34 66.37 14.1 33.5 34.05 34.5 495.5526681
97 34.5 67.83 2.92 34 34.05 35 500.5373567 35 87.92 40.18 34.5 34.05 35.5 509.7423599 35.5 105.47 35.1 35 34.05 36 520.8035776 36 134.33 57.72 35.5 34.05 36.5 527.7145947 36.5 134.47 0.28 36 38.57 37 533.2632824 37 144.71 20.48 36.5 38.57 37.5 538.5456181 37.5 144.71 0 37 38.57 38 546.1310538 38 144.71 0 37.5 38.57 38.5 552.5431178 38.5 144.71 0 38 38.57 39 562.9381711 39 145.5 1.58 38.5 38.57 39.5 573.211262 39.5 145.5 0 39 38.57 40 581.85678 40 145.9 0.8 39.5 42.79 40.5 149.49 7.18 40 47.36 41 153.22 7.46 40.5 59.60 41.5 158.67 10.9 41 76.09 42 171.71 26.08 41.5 89.48 42.5 183.81 24.2 42 112.33 43 196.98 26.34 42.5 133.87 43.5 211.47 28.98 43 134.85 44 226.1 29.26 43.5 137.30 44.5 243.39 34.58 44 137.95 45 266.27 45.76 44.5 137.95 45.5 278.11 23.68 45 143.50 46 291.41 26.6 45.5 147.75 46.5 304.57 26.32 46 147.75 47 325.32 41.5 46.5 166.35 47.5 345.27 39.9 47 172.06 47.5 174.35 48 189.54 48.5 230.99 49 258.41 49.5 258.57 50 259.55 50.5 259.88 51 259.88 51.5 260.53 52 272.61
98 52.5 276.70 53 288.28 53.5 304.94 54 325.01 54.5 337.58 55 347.69 55.5 357.33 56 371.20 56.5 382.95 57 402.05 57.5 420.99 58 436.98 58.5 453.71 59 470.67 59.5 486.63 60 504.59 Data for Figure 4.4
0 0 0 0 0 0 0 0.0% 0.0% 8 0.156 0.032 12 0.105 0.02 24 12.0% 3.4% 16 0.707 0.03 24 0.161 0.031 36 72.5% 2.2% 18 0.764 0.04 35 0.801 0.03 48 77.4% 2.6% 20 0.739 0.02 42 0.765 0.03 56 72.9% 4.2% 23 0.724 0.05 47 0.809 0.035 58 73.9% 3.9% mean 0.792 79.2 60 73.5% 4.9% SD 0.023 2.3 62 75.3% 3.4%
0 0.051 0.02 0 0 0 10 0.154 0.02 14 0.085 0.02 16 0.536 0.01 24 0.246 0.01 20 0.732 0.02 26 0.56 0.024 22 0.746 0.03 28 0.69 0.025 24 0.727 0.01 30 0.71 0.026 26 0.725 0.02 32 0.71 0.031 28 0.719 0.01 40 0.7 0.019 30 0.734 0.005 32 0.741 0.01
99 Data for Figure 4.5
Glu In Glu Eff 0 3289 154 3321 162 10 3354 158 1874 134 16 3319 152 1053 131 20 3284 146 863 120 24 3249 70 849 114 28 3214 164 841 116 32 3279 148 861 121
Acetate Butyrate 0 0 0 0 0 10 243 18.5 74 3 16 582 24.3 195 11 20 747 32 336 15 24 843 30 324 12 28 798 38 342 16 32 822 34 321 14
100 Husen Zhang
Education 2005 Ph.D. Environmental Engineering, The Pennsylvania State University 2000 M.S., Environmental Engineering, The Pennsylvania State University 1998 B.S., Environmental Engineering, Tsinghua University
Experience Aug 1998 – Dec 2004 Research Assistant, The Pennsylvania State University Jan 2001 – Jan 2004 Graduate Fellow, Biogeochemical Research Initiative for Education
Awards
2000 Second-Place Oral Presentation Award, Zhang H and BE Logan. 2000. Biodegradation kinetics of perchlorate reducing bacteria. American Society for Microbiology Allegheny Branch, Fall Meeting, Environmental Microbiology Session. Oct 27-28. 1998 Graduate Dean's Fellowship, The Pennsylvania State University.
Publications
Zhang H, BE Logan, JM Regan, LA Achenbach, and MA Bruns. 2004. Molecular assessment of inoculated and indigenous bacteria in biofilms from a pilot-scale perchlorate-reducing bioreactor. Microbial Ecology, In press. DOI: 10.1007/s00248-004-0273-6
Iyer, P, Bruns MA, Zhang H, vanGinkel S, and BE Logan. 2004. H2-Producing bacterial communities from a heat-treated soil inoculum. Applied Microbiology and Biotechnology. 66(2): 166-173
Zhang H, Bruns MA, and BE Logan. 2002. Perchlorate reduction by a novel chemolithoautotrophic, hydrogen-oxidizing bacterium. Environmental Microbiology, 4(10): 570-576.
Logan BE, H Zhang, P Mulvaney, MG Milner, IM Head, and RF Unz. 2001. Kinetics of perchlorate- and chlorate- respiring bacteria. Applied and Environmental Microbiology, 67(6): 2499-2506.
Wu J, RF Unz, H Zhang and BE Logan. 2001. Persistence of perchlorate and the relative numbers of perchlorate- and chlorate-respiring microorganisms in natural waters, soils and wastewater. Bioremediation Journal, 5(2):119-130.
Manuscripts in Preparation
Zhang H, Bruns MA, and BE Logan. 2004. Biological hydrogen production from an unsaturated-flow, packed-bed column reactor. For submission to Water Research