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ADVANCES IN

Applied Microbiology VOLUME 38

This Page Intentionally Left Blank

ADVANCES IN

Applied Microbiology Edited by SAUL NEIDLEMAN Vacaville, California

ALLEN 1. LASKIN Somerset, New Jersey

VOLUME 38

Academic Press, Inc. Harcourt Brace Jovanovich, Publishers

San Diego New York Boston London Sydney Tokyo Toronto

This book is printed on acid-free paper. @

Copyright 0 1993 by ACADEMIC PRESS, INC. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher.

Academic Press, Inc. 1250 Sixth Avenue, San Diego, California 92101-431 1

United Kingdom Edition published by

Academic Press Limited 24-28 Oval Road, London NWI 7DX

Library of Congress Catalog Number: 59-13823 Inteniational Standard Book Number: 0-12-002638-4 PRINTED IN THE UNITED STATES OF AMERICA 93 94 95 96 97 98 BB 9 8 1 6 5 4 3 2 1

CONTENTS

Selected Methods for the Detection and Assessment of Ecological Effects Resulting from the Release of Genetically Engineered Microorganisms to the Terrestrial Environment

.

.

G STOTZKY. M. W BRODER.J. D . DOYLE.AND R . A . JONES I. Introduction ......................................................... I1. Methods of Study .................................................... 111. Representative Results ................................................ IV. Discussion ........................................................... V. Summary ............................................................ References ...........................................................

2 7 50 90 93 95

Biochemical EngineeringAspects of Solid-state Fermentation

M. V. RAMANAMURTHY.N . G . KARANTH. AND K . S. M . S. RAGHAVARAO I . Introduction ......................................................... I1. Mass Transfer in Solid-state Fermentation Systems ......................

. Heat Transfer in Solid-state Fermentation Systems ...................... . Influence of Bioreactor Design on Mass Transfer ......................... Heat Dissipation in Solid-State Fermentation Bioreactors ................. . Role of Water Activity ................................................ . Important Physical Parameters in Solid-state Fermentation ............... . Mathematical Modeling in Solid-state Fermentation Systems .............

111 IV V. VI VII VIII

IX. Experimental Measurements ..........................................

X . Conclusions ......................................................... XI . Nomenclature ........................................................ References ...........................................................

99

102 106 108 111 112 114 118 130 141 142 144

The New Antibody Technologies

ERIK P. LILLEHOJAND VEDPAL s. MALIK I. A Brief History ....................................................... I1. Polyclonal Antibodies ................................................

111. Immunoglobulin-Binding Proteins from Bacteria ........................ IV. Recent Developments in Antibody Purification .......................... V. Monoclonal Antibodies ............................................... VI . Recombinant Antibodies ..............................................

V

150 151 157 158 161 170

vi

CONTENTS

. . . . .

VII Antibody Immunotherapy ............................................. VIII . Immunotoxins ....................................................... IX Radiolabeled Antibodies in Clinical Medicine ........................... X Immunofluorescent and Immunomagnetic Techniques ................... XI . New Enzyme Immunoassay Techniques ................................ XI1 Antibody Uses in Recombinant DNA Technology ........................ XI11 Antibodies in the Future .............................................. References ...........................................................

180 182 186 186 187 191 194 195

Anoxygenic Phototrophic Bacteria: Physiology and Advances in Hydrogen Production Technology

. U G H U V E E R RAO. AND K . L. KOVACS

K . SASIKALA. CH. v. U M A N A . P

.

I Introduction ......................................................... I1. Classification and Growth ............................................. 111. Methods Used for Hydrogen Metabolism Studies ........................ IV Production of Hydrogen by Purple Nonsulfur Anoxygenic Phototrophic Bacteria ............................................................. V. Enzymes Related to Hydrogen Metabolism .............................. VI . Carbon Assimilation .................................................. VII . Advances in Hydrogen Production Technologies Using Anoxygenic Phototrophic Bacteria ................................................. VIII Other Uses of Anoxygenic Phototrophic Bacteria ........................ IX Conclusion .......................................................... References ...........................................................

.

. .

INDEX.................................................................... CONTENTS OF PREVIOUS VOLUMES ..........................................

211 213 219 220 240 259 267 279 281 281

297 316

Selected Methods for the Detection and Assessment of Ecological Effects Resulting from the Release of Genetically Engineered Microorganismsto the Terrestrial Environment G. STOTZKY,M. W.BRODER,'J. D.DOYLE,~ AND R. A. JONES3 Laboratory of Microbial Ecology Department of Biology New York University New York, New York 10003

I. Introduction 11. Methods of Study A. Soil Preparation B. Metabolic Activity C. Preparation of Soil for Enzyme and Microbial Assays D. Microbial Assays E. Soil Enzymes F. Nitrogen Transformations G. Nonsymbiotic Dinitrogen Fixation H. Growth Rates and Competitive Ability of Genetically Engineered Microorganisms I. Statistical Design and Analysis 111. Representative Results A. Metabolic Activity (Carbon Dioxide Evolution) B. Species Diversity C. Activity of Soil Enzymes D. pH E. Effect of Adding 2,4-Dichlorophenoxyacetateand a Genetically Engineered Microorganism Capable of Its Catabolism on Microbial Populations and Processes in Soil F. Nitrogen Transformations G. Survival of Genetically Engineered Microorganisms and Their Homologous Parents in Soil IV. Discussion V. Summary References

'Present address: U.S. Environmental Protection Agency, Washington, D.C., 20460. *Present address: ManTech Environmental Technology, Inc., U.S. Environmental Protection Agency, Corvallis, Oregon 97333. 3Present address: Food and Drug Administration, Rockville, Maryland 20855.

1 ADVANCES IN APPLIED MICROBIOLOGY,VOLUME 38 Copyright 8 1993 by Academic Press, Inc. All rights of reproduction in any form reserved.

2

G.STOTZKY ET AL.

1. Introduction

The use of microorganisms as alternatives to traditional chemical and physical technologies is being explored in such areas as agriculture, pest control, and bioremediation of toxic wastes. These applications of biotechnology rely on the expression of useful genetic traits in both naturally occurring microorganisms and microorganisms genetically modified by recombinant DNA techniques. In the latter, the merging of the fields of molecular biology and microbial ecology is providing exciting alternative technologies, as well as new uncertainties. These uncertainties are associated with (1)the environmental uses of genetically engineered microorganisms (GEMs) capable of expressing traits not present in the unmodified parent microorganism; (2) the probability of the transfer of these genetic traits to other microorganisms indigenous to the environment; and (3) the possibility of the new traits having a deleterious effect on the environment. The potential risks to public health and the environment from a deliberate or accidental release of GEMs to the environment are the most urgent concerns, both scientifically and with respect to public policy, associated with this aspect of biotechnology. Questions about the probabilities of survival, colonization, and function of released GEMs and their novel DNA in natural habitats and the ability to predict the consequences of their release will be answered only by applying the knowledge derived from the study of the ecology and molecular interactions among microbes in these habitats. Both biotic and abiotic environmental characteristics affect the survival, perpetuation, efficacy, and risk associated with the release of novel DNA in GEMs to any natural habitat. Moreover, the survival of novel genetic information and its potential effects on the homeostasis of an ecosystem may be greater if the information is transferred to indigenous species that are more adapted to the specific habitat than the introduced GEMs. The purpose of this article is to summarize the methods and concepts developed and used by the authors to study the potential effects of GEMs on microbial populations and microbe-mediated ecological processes in soil. The potential impacts of GEMs, unrelated to the purposes for which they were engineered, on the structure and function of the natural environments into which they are introduced constitute the bottom-line concern about the release of GEMs to the environment. If a GEM survives in the habitat into which it is introduced and does the job for which it was designed, and even if the novel gene(s) is transferred to indigenous microorganisms, there should be little cause for concern unless the novel gene(s), either in the introduced GEM or in an

RELEASE OF GENETICALLY ENGINEXRED MICROORGANISMS

3

indigenous recipient(s), results in some unexpected impacts on the environment. This concept is easy to state but difficult to translate into an effective experimental design. What effects [i.e., environmental perturbations) should be sought, especially if the novel gene(s) codes for a limited function(s) and the GEM has been selected or programmed for poor survival in a specific habitat? Considering the current state of the art and the paucity of data on detection, enumeration, survival, growth, and transfer of genetic information [both intra- and interspecifically) by microorganisms in natural habitats, the detection, measurement, and evaluation of potential effects of an introduced GEM on ecological processes is akin to finding a needle in a haystack. However, this is the most pertinent concern about the release of GEMs to the environment, and more studies on this aspect must be conducted. Nevertheless, as insufficient basic knowledge is available about the fate of introduced microorganisms, whether genetically engineered or not, in natural habitats, data from studies, especially in microcosms rather than in the field, on the ecological effects of GEMS must be interpreted and applied cautiously to avoid establishing far-reaching and long-lasting policies, criteria, and regulations that may be based on incomplete or erroneous data. The microbe-mediated ecological processes that should be evaluated before the release of a GEM to the environment should be those for which techniques are well established, that cover a broad spectrum of relevant microbial activities, and that have been successfully used to study the perturbation of the soil environment by chemical and physical factors: for example, (1)metabolic activity and carbon mineralization, as measured by CO, evolution or other respiratory techniques; (2) transformations of fixed nitrogen by perfusion techniques; (3) dinitrogen fixation, using the acetylene-reduction technique; (4)species diversity of the microbiota, using selective and differential media; and (5) activity of selected enzymes, such as acid and alkaline phosphatases [to provide a measure of the cycling of P), arylsulfatases [to provide a measure of the cycling of S ) , and dehydrogenases (to provide another measure of overall metabolic activity). These processes should be monitored for extended periods after the introduction of a GEM, whose fate, as well as that of its novel gene(s), should be concurrently followed. Some desirable characteristics of methods for assessing the ecological effects of GEMs are presented in Table I. In addition to evaluating the potential effects of GEMs on these defined ecological processes, the investigator should be alert to the possible occurrence of unanticipated effects that cannot be predicted from

4

G. STOTZKY ET AL. TABLE I DESIRABLE CHARACTERISTICS OF METHODS FOR ASSESSING ECOLOGICAL EFFECTSOF GENETICALLYENGINEERED MICROORGANISMS Relevance Representative of the microbial community Sensitivity Reproducibility Ease (facility; rapidity) Cost-effectiveness Interlaboratory validation Predictiveness (transferability;modeling) Ecological versus statistical significance

the information encoded on the novel DNA (i.e., pleiotropic effects). For example, the acquisition of a plasmid carrying genes for dinitrogen fixation and antibiotic resistance by various species of phytopathogenic bacteria apparently resulted in a spectrum of unrelated and unpredicted biochemical and physiological alterations (Kozyrovskaya et al., 1984). Other pleiotropic effects resulting from the insertion of novel genes have been reported (e.g., Stotzky, 1989; Stotzky and Babich, 1986).If pleiotropic effects are indicated, the battery of tests for ecological effects should be extended, as such unanticipated alterations could affect ecological processes in soil and other environments. Furthermore, the growth rates of the GEMs, as well as of the homologous microorganisms without the novel gene, and their ability to compete with indigenous microbes in soil should be determined (e.g., by the soil replica-plating technique). The purpose and function of the introduced novel genes must be considered in the design of the studies. For example, if a GEM carries a novel gene(s) that codes for a catabolic function (e.g., the degradation of a xenobiotic), the soil should be amended with the appropriate “substrate” on which the products of the novel gene(s) act, to determine whether the gene provides ecological advantages to the GEM, whether intermediates are produced, and whether and how these advantages and intermediates affect ecological processes. When GEMs resistant to the toxicity of heavy metals or other antimicrobial agents are used, the soil should be stressed with the specific agents to which the novel genes confer resistance. When GEMs containing nif genes are used, nitrification rates and attendant decreases in pH should be monitored, as enhanced dinitrogen fixation could increase nitrification and the accumulation of protons, which could affect numerous ecological processes.

RELEASE OF GENETICALLY ENGINEERED MICROORGANISMS

5

To verify that the novel gene(s) is responsible for any changes in the ecological processes evaluated, all studies should be conducted in parallel in the same soils inoculated with equal amounts of the GEM or the homologous microorganism without the novel gene(s). The existing numbers of the introduced GEM [or its novel gene(s) in another recipient] should be related, over time, to the magnitude of perturbation of any of the ecological processes. The duration of these effects should also be determined, especially after the GEM or its novel gene(s) can no longer be detected. These studies should be conducted concurrently in several laboratories, to obtain interlaboratory validation and to enhance the development of appropriate procedures with which to study the effects of GEMs on ecological processes in soil. A major goal of the studies should be the development of a standard battery of assays that will most clearly, rapidly, easily, and inexpensively detect any ecological effect of introduced GEMs and that can be used in the assessment of the risk of introducing any GEMs, either purposely or inadvertently, into soil and other natural habitats. The studies should be conducted initially in the laboratory, because the potential risks associated with the introduction of a GEM to the environment are unknown. A variety of terrestrial microcosms that purportedly simulate field conditions have been developed. These microcosms range from extremely simple systems that inoculate a GEM into sterile soil added to a sterile nutrient broth in test tubes (e.g., Walter et a]., 1987);to sterile or nonsterile soil in a test tube, flask, or other container (e.g., Stotzky et al., 1990);to multiple containers of nonsterile soil enclosed within a larger container (e.g., Fig. 1); to more complex systems that involve undisturbed soil cores of varying sizes brought into the laboratory with minimum disturbance of the structure and biotic composition of the soil (e.g., Bentjen et al., 1989;Fredrickson et al., 1989;Hicks et al., 1990;Van Voris, 1988); to either disturbed or undisturbed soils that are cropped and maintained within chambers that enable the control of temperature, relative humidity, light/dark cycles, and other environmental variables (Fig. 2) (e.g., Armstrong et al., 1987;Gile et al., 1982;Knudsen et al., 1988). Examples of microcosms with different degrees of complexity and the rationales for their use have been discussed (e.g., Atlas and Bartha, 1981;Cavalieri, 1991;Gillett, 1988;Greenberg et al., 1988;Hicks et a]., 1990;Johnson and Curl, 1972;Pritchard, 1988; Pritchard and Bourquin, 1984; Stotzky et al., 1990). Guidelines for the use of soil core microcosms, with descriptions of various core designs, sampling procedures, and statistical analyses, have been published (Van Voris, 1988). The microcosms and techniques used by the authors to study the

FIG. 1. Incubation unit for measuring CO, evolution from soil. The unit is used when subsamples of soil are to be removed during incubation. When this is not required, soil is placed directly into the master jar, which can then be smaller (Stotzky,1965a).


7

Air inlet Rain

FIG. 2. Microcosm in which environmental variables (e.g., temperature, relative humidity, lightldark cycles] can be controlled (Gile et al., 1982).

effects of GEMs on some microbial populations and processes in soil are described herein. Although other microcosms and techniques are available (e.g., Page et al., 1982; Nannipieri et al., 1990; and as referenced above), only procedures with which the authors have hands-on experience with GEMs in soil are discussed. II. Methods of Study

A. SOILPREPARATION Sieve soil (top -5 cm) collected in the field through a broad-mesh screen (e.g., 1 cm) to remove stones and plant debris and to disrupt large soil aggregates. Mix the sieved soil thoroughly to provide as uniform and representative a sample as possible. The sieved soil can be

8

G.STOTZKY ET AL.

used immediately after collection or it can be stored. Although soil used immediately probably better reflects the microbiological conditions that exist in the soil in situ, there are disadvantages: for example, if the same soil is to be used in subsequent experiments, collection from adjacent sites and in different seasons can result in both biotic and abiotic variability. Moreover, if the desired soil is located some distance from the laboratory, considerable time will be consumed in collection. The collection, sieving, and storage of quantities of soil from the same site sufficient for numerous experiments eliminate these disadvantages. Changes in the microbiota as the result of storage of the soil can be rectified to some extent (Stotzky et al., 1962). For example, soils can be maintained in wooden flats (e.g., 55 x 30 x 15 cm) in a greenhouse under a regime of intermittent cropping and fluctuating temperatures and moisture (Stotzky, 1973). If a greenhouse is not available, soils can be stored at room temperature in large plastic or metal garbage cans lined with plastic garbage bags. Two weeks before the initiation of a study, pass the soil through a 2-mm sieve, and rejuvenate the soil by bringing it to - 33-kPa water tension and adding glucose (I%,w/w, in a mineral salts solution) and approximately 20 mg of fresh garden soil per gram soil, oven-dry equivalent. Maintain the soil at room temperature, and mix every few days (Devanas et a]., 1986). When soil is to be amended with clay minerals, mix the soil after the initial sieving with the appropriate mined clay mineral. Use an electricpowered cement mixer for uniform and rapid mixing. The desired ratios of clay and soil can be achieved on a weightjweight or a volume/ volume basis, although the latter (using buckets) is more conveniently conducted in the field, especially with large volumes of soil. B. METABOLIC ACTIVITY

The overall metabolic activity of microbes in soil can be determined with respirometric techniques that monitor either CO, evolution or 0, consumption. These methods, especially when CO, evolution is measured, probably provide the best and most easily measured index of the gross metabolic activity of mixed microbial populations in soil (Anderson, 1982; Stotzky, l960,1965a, 1972). The “master jar” system (Fig. 1) (Stotzky, 1965a; Stotzky et al., 1958) enables the removal of subsamples of soil during an extended incubation for various analyses (e.g., transformation of substrates, species diversity, enzyme activities, survival of introduced microorganisms, including the GEMS and their novel genes) without disturbing the remaining soil. Sampling without disturbance


9

eliminates artifactual peaks in CO, evolution resulting from the physical disturbance of the soil (Stotzky and Norman, 1961a,b, 1964). The soils are incubated at controlled temperatures and maintained at their - 33-kPa water tension by continuous aeration with water-saturated, C0,-free air. The amount of CO, trapped in NaOH collectors is determined, after precipitation of the CO, with BaCl,, by automatic potentiometric titration with HC1. The amount of CO, evolved from the master jars during an incubation is expressed on the basis of a constant amount of soil, usually 100 g, oven-dry equivalent, which normalizes the respiration rate regardless of the amount of soil present in the master jars. The potential gross metabolic activity, both aerobic and anaerobic, of the heterotrophic soil microbiota can be measured by the addition of a nonspecific substrate (e.g., glucose), and the potential activity of specific populations can be evaluated by the addition of specific substrates (e.g., celluloses, starches, lipids, proteins) whose mineralization is dependent on the ability of these populations to synthesize the appropriate enzymes. In particular, aldehydes, which are highly selective substrates, can be used (Bewley and Stotzky, 1984; Kunc and Stotzky, 1974, 1977). Ratios of the gross metabolic activity (with glucose or other nonspecific substrates) to that of specific metabolic activities (e.g., with aldehydes or other selective substrates) can be used to indicate whether the presence of a GEM exerts an effect on the metabolism of all components of the indigenous microbial population or only on certain segments of the microbiota. These ratios will also sharpen comparisons between uninoculated control soils and soils inoculated with either a GEM or the homologous parental strain without the novel gene. When aldehydes are used as specific substrates, the data from the aldehyde studies should be correlated with those from nitrification studies. Both nitrification (an autotrophic process) and mineralization of aldehydes (a heterotrophic process) are restricted to certain but different microbial species, and both processes show similar kinetics in soil, especially when soils are stressed (e.g., with heavy metals or acid precipitation) or altered (e.g., amended with different clay minerals) (Stotzky, 1980,1986). The soils should also be amended with the specific substrate (e.g., toluene, xylenes, 2,4-dichlorophenoxyacetate)on which the products of the novel gene(s) in a GEM function, to determine whether the substrate provides an ecological advantage to the GEM, whether intermediates are produced from the substrate, and how any advantages or intermediates affect both nonspecific and specific metabolic activities, as well as other microbe-mediated ecological processes (Doyle et a ] . ,

10

G . STOTZKY ET AL.

1991; Short et al., 1991). When the GEM contains genes that confer resistance to the toxicity of an antimicrobial agent, the soil should be amended with the appropriate agent, to determine whether such a stress (simulated worst-case scenario) confers an ecological advantage on the GEM and whether this advantage, in turn, influences the activity and population dynamics of the indigenous microbiota. Measurement of Respiration [Carbon Dioxide Evolution) Reagents NaOH (-1.5 N): Dissolve, with swirling, approximately 60 g NaOH per liter of distilled water in a 20-liter borosilicate carboy. Fit into the mouth of the carboy a two-hole rubber stopper containing an air inlet tube and a solution outlet tube that is attached to a constantvolume 50-ml stopcock-type automatic pipetter. Connect the air inlet tube to a gas-drying tube containing Drierite and Ascarite, to prevent ambient water vapor and CO, from entering the NaOH, and attach a rubber bulb to enable pressurization of the carboy. During the course of the study, adjust the normality of the solution to the amount of CO, produced. NaOH (2.00 N standard): Commercially available. HC1 (-7.5 N): Dilute approximately 625 ml of concentrated HC1 to 1 liter with distilled water (add the HC1 to the water). As the normality of the NaOH in the CO, collector is adjusted to reflect decreasing or increasing (e.g., following pulsing with a carbon source) respiration rates, the normality of the HC1 must also be changed to reflect the 5-fold difference between the normality of the NaOH and that of the HC1. (The CO, collector contains 50 ml of NaOH, and the self-filling burette on the automatic titrator has a capacity of only 10 ml, to enhance precision.) Determine accurately the normality of the HC1 by titrating it against the 2.00 N NaOH standard. Attach a Drierite-Ascarite column and rubber bulb to the air inlet tube, as described for the NaOH carboy. Attach the solution outlet tube to the self-filling burette. BaC1, (-3.5 M): Dissolve approximately 855 g BaC1,-2H,O per liter of distilled water. Place the solution in a glass carboy equipped with a Drierite-Ascarite column and rubber bulb on the air inlet tube, and connect the solution outlet tube to a 50- or 100-ml selffilling burette. Distilled water: Fill a glass carboy with freshly distilled water. Attach a Drierite-Ascarite column and rubber bulb to the air inlet tube and an eyedropper tube to the outlet tube. The flow of water is controlled with a pinch clamp on the outlet tubing.


11

KOH (-4.0 N): Dissolve approximately 224 g KOH per liter of distilled water in either a borosilicate flask or carboy. Transfer the solution to the appropriate glass carboys in the scrubber system of the respiration train. (N.B.: The dissolution of NaOH and KOH is an exothermic reaction and requires the use of a borosilicate container. The solutions must be allowed to cool before transferring to other containers that are not heattolerant.) Procedure. Grow the GEMs and the homologous parental strains as batch cultures to a population of around loQ colony-forming units (CFU)per milliliter in an appropriate liquid medium (e.g., L-broth) containing the selection factors necessary to maintain the genotypic and phenotypic characteristics unique to the GEMs and the parental strains and necessary for their selective recovery from soil. Prepare a standard curve for each GEM and homologous parental strain by plotting the absorbance against the numbers of either total (determined microscopically, e.g., with a hemacytometer) or viable bacteria (determined by plating) for a dilution series. Determine the concentration of the bacteria spectrophotometrically (e.g., with a Bausch & Lomb Spectronic 20) at the same wavelength used for the preparation of the standard curve, using sterile medium as a blank. Dilute the bacteria with a sterile substrate (e.g., glucose) solution or sterile water and add to the soil with sufficient water to adjust the soil water tension to - 33 kPa and to yield the desired inoculum density per gram of soil, oven-dry equivalent (e.g., one that approximates the density to be used in a field release), and the desired substrate concentration. Mix the soil in a thin-walled plastic bag by kneading, and store for 48 hours at 4"C, with additional kneading at 24 hours, to enhance the uniform distribution of water, substrate, and cells. After the 48-hour equilibration period, weigh 50 g of soil, oven-dry equivalent, into 100cm3 glass vials. If cold-intolerant microorganisms are used, add the cells just before weighing the soils, and mix well. Keep the vials of soil cool until all have been filled. Place the vials into a wide-mouth gallon jar (master jar) (Fig. 1)(pickle and mayonnaise jars are ideal and can be obtained inexpensively). Attach the master jar, via the air inlet tube, to the manifold of a respiration train that contains a scrubber system for removing oil, ambient COz, various nitrogen compounds, and other contaminants and then resaturates with water the air that continuously flushes the master jar (Fig. 3). Connect the air outlet tube of the master jar to the CO, collector (Fig. 1) (see below for details). At specified in-

12

G.STOTZKY ET AL.

4

I KOH

4

&

KOH

FIG. 3. Schematic of scrubber system used to remove contaminants and C 0 2from and to saturate with water the air that flushes the master jars.

tervals, remove a soil vial from each master jar for microbiological, enzymatic, physical, and chemical analyses (Fig. 4). Immediately after placing the soil vials into the master jars, analyze the soil (most efficiently done with soil remaining after the vials have been filled) for the microbiological, enzymatic, physical, and chemical characteristics that will be analyzed during the experiments. These constitute the data for day 0. Flush the soils in the master jars continuously with C0,-free, watersaturated air, to remove respired CO, and to maintain the soils at the - 33-kPa water tension. Respiration rates are determined by trapping the evolved CO, in NaOH and periodically titrating the unneutralized NaOH with HCl, contained in a 10-ml self-filling burette, with an automatic titrator (e.g., Radiometer TTTBO) connected to a pH meter (e.g., Radiometer PHM82 standard pH meter). Place into each master jar sufficient soil vials for the number of subsamples to be analyzed during the course of the study (it is advisable to place more soil vials than needed, in the event of any contingencies, such as dropping a vial or the need to continue the experiment longer than originally designed). Insert the rubber stopper (No. 15), with the air inlet and outlet tubes, and secure it with a wire spring, the ends of


13

' I Master jar

Subsample

m2Evolution

Soil enzyme asap

Sebcliw media I C3ped.r divercity

NUlritiod

orow

Antibiogrmr

I Totd m e r i a

I

GEM media

CHO Utilization/ Fermentation

I

Biochemical idec4ification

Physical chwaderization

I

DNA pmbe

Gel

eledmphoreeis

Restridion enzyme mwiq

FIG.4. Flowchart of microbiological and enzymatic analyses conducted on subsamples of soil from the master jar (see Fig. 1).

which are inserted into loops twisted in a wire circle that is fastened around the neck of the jar (Fig. 1).Attach the master jar, via the air inlet tube, to the manifold of the respiration train. The air that flushes the master jars must be treated to remove inorganic and organic contaminants and ambient CO, and then saturated with water. The air scrubber system (Fig. 3) consists of (1)a pressure regulator with coarse adjustment; (2) two filters (e.g., HI5J-6C10-025 and HI5J-AU10-025, Finite Filter, Oxford, MI) in series, to remove particulates, oil droplets, and other contaminants; (3) a pressure regulator with fine adjustment (pressure range 0 to 25 psi]; (4) a shut-off valve; (5) a water-filled manometer constructed of Tygon tubing [e.g., 13 mm inner diameter (ID)]that extends from the floor to the ceiling, is open to the atmosphere, and is vented to a sink or flask (this manometer serves as a pressure-release valve in the event of a blockage in the airflow system; a water column of -0.7 m is equivalent to 1 psi); (6) an empty 20-liter glass carboy that serves as a trap; (7) two 20-liter glass carboys, each containing 5 liters of 4.0 N KOH (replace the KOH every 2 weeks, as K,CO, will eventually form in the bottom of the air inlet tube, disrupting the air flow, and the KOH will eventually be neutralized by the CO,; to prevent disruption of the air flow during replacement, two pairs of 20-liter carboys, each containing 5 liters of 4.0 N KOH, are placed in parallel, and the air flow is shunted between the carboys by the appropriate placement of pinch clamps]; ( 8 ) a second empty trap; (9) two 20-liter glass carboys, each containing 6 liters of

14

G. STOTZKY ET AL.

distilled water to rehumidify the air (replace the water periodically, as it may become contaminated with KOH); and (10)a flow meter. The components of the scrubber system are connected with glass or plastic, straight or T-shaped, tubing connectors or plastic one-way valves [6.4 mm outer diameter (OD)]and latex tubing (6.4 mm ID). Each carboy has air inlet and outlet tubes attached through a rubber stopper in the mouth of the carboy. The air inlet tube extends 4 to 6 cm into the solution, and the outlet tube protrudes slightly below the rubber stopper. In the traps, the air inlet tube protrudes slightly below the stopper, and the outlet tube extends almost to the bottom of the carboy. This arrangement of inlet and outlet tubes in the traps enables the KOH and water to be pushed back into their carboys from the traps in the event that the airstream is disrupted and the one-way valves malfunction. The air scrubber system is attached to a manifold that distributes the C0,-free, water-saturated air to each master jar. The manifold consists of a series of glass or plastic T-shaped tubing connectors (6.4 mm OD), the side arms of which are connected to adjacent T-shaped tubing connectors with latex tubing (6.4 mm ID, 1.6 mm wall thickness), and the perpendicular arm is attached to the air inlet tube of the master jar. A 25-gauge hypodermic needle, which serves to equalize the air flow to each master jar, is inserted into the perpendicular arm of the T-shaped tubing connector and is fixed in place by the latex tubing that covers the perpendicular arm at one end and extends to one-half of a plastic quick-disconnect connector at the other end. A one-way valve, to prevent back flow of air with a loss of CO, from the master jar in the event of a reduction in pressure in the air-flow system, is inserted in this tubing. The other half of the quick-disconnect connector is inserted into the end of the tubing that is attached to the air inlet tube of the master jar. The master jar is closed with a No. 15 rubber stopper that contains two glass tubes, both of which are attached to quick-disconnect connectors: one glass tube is the air inlet and terminates on the underside of the stopper; the other tube, which extends to the bottom of the master jar to prevent channeling of the airstream, is the air outlet. The manifold tubing (containing a quick-disconnect connector) is connected to the air inlet of the master jar, and the outlet tubing of the master jar (containing a quick-disconnect connector) is connected to the air inlet of the CO, collector. The CO, collector is a 200-cm3 glass tumbler closed with a rubber stopper (No. 13%) in which an adjustable glass column (chimney) (300 x 17.5 mm), which extends into the NaOH in the tumbler, and a


15

glass air inlet tube are inserted (Fig. 1).The base of the chimney is constricted to retain glass beads (6 mm), which disrupt air bubbles and increase the surface area of the NaOH, thereby maximizing the absorption of CO,. The C0,-free air is then vented to the atmosphere. Studies with 14C-labeledsubstrates have shown that the absorption of CO, in the collectors is 100% efficient (Stotzky, 1965a). Before connecting the master jars to the manifold, set the air pressure gauge to about 3 psi and open the air valve. Connect the master jars to the manifold, and close with screw clamps any outlets on the manifold that are not connected to a master jar. Include several empty master jars, interspersed among the jars containing soil, to serve as blanks for nonrespired CO,. Purge the master jars for approximately 30 min with C0,free air. Fill and empty the 50-ml stopcock-type automatic pipetter four times with the NaOH solution to remove old NaOH and any precipitate. Fill the pipetter completely (50 ml) with NaOH, and dispense the NaOH into a glass tumbler. Insert the stopper with the chimney, and connect the air inlet tubing to the outlet tubing of the master jar via the quick-disconnect connector. The continuous airstream will force the NaOH into the chimney. When all CO, collectors are connected, adjust the air flow through each master jar to comparable rates by raising or lowering the chimneys. Keep the outside of the chimneys in the vicinity of the stoppers lubricated with silicone grease, and wear a heavy glove when adjusting them (the glass chimneys become brittle after ex'tensive use and can break during adjustment). Adjust the rate of air flow to 10 to 15 literdhow; monitor the air flow with the flow meter. The titration schedule will depend on when the substrate was added, the nature of the substrate, and the type of data desired. With glucose as the substrate, titrations are usually conducted daily for the first 5 to 7 days, then on alternate days for the subsequent week, and then on every third or fourth day for the remainder of the study. To determine the amount of CO, evolved, disconnect the tumbler from the master jar at the quick-disconnect connector, loosen the rubber stopper, and rinse the glass beads and the inside of the chimney, as well as the outside of the bottom of the chimney, with about 100 ml of distilled water into the tumbler. Add approximately 10 ml of the BaC1, solution to the tumbler to precipitate the adsorbed CO, as BaCO,. (The amount of BaC1, added depends on the amount of CO, evolved. Add BaC1, until no more precipitate is formed.) Place the tumbler on a magnetic stir plate, add a magnetic stir bar, and insert the pH electrodes (glass and calomel or a combination electrode) and the capillary HC1-delivery tube attached to the magnetic valve of the automatic titrator. Fill the burette with HC1,

16

G. STOTZKY ET AL.

and adjust the capillary tube so that it dispenses the HCl onto the glass electrode. Turn on the magnetic stir plate, and start the titration. When the titration is complete, read on the burette the amount of HC1 required to neutralize (pH 7.0) the NaOH in the tumbler and record. Discard the neutralized NaOH, wash the tumbler with tap water, rinse with distilled water, refill with 50 ml of NaOH, insert a stopper with the chimney, and reattach the CO, collector to a master jar. Immediately after removing the CO, collector for titration and when not removing samples of soil, replace it with a fresh collector. Filling a tumbler from one master jar with fresh NaOH, washing the beads and chimney and adding BaC1, to a collector to be titrated, discarding neutralized NaOH, and washing the tumbler are done while a tumbler from another master jar is being titrated. Do not pour the neutralized NaOH down a sink drain, but pour into a large bucket or carboy. Allow the BaCO, to accumulate on the bottom, decant the clear supernatant, and dispose of the concentrated BaCO, according to regulations for the disposal of hazardous wastes. When samples of soil are removed from the master jar after titration, purge the ambient air from the master jar for about 30 min before attaching a CO, collector. Calculate the amount of carbon (C) respired by using the formula mgC

=

(B

-

S)6N

where B is the average amount (ml) of HC1 required to neutralize the NaOH in the CO, collectors attached to empty master jars (blanks), S is the amount (ml) of HCl required to neutralize the NaOH in the sample CO, collector, 6 is the equivalent weight of C, and N is the normality of the HC1 solution. OF SOILFOR ENZYMEAND MICROBIAL ASSAYS C. PREPARATION

Periodically remove a soil vial from each of the master jars, and subject the soil to a variety of microbial, enzymatic, and other analyses (Fig. 4). Weigh approximately 10 g of soil from the vial into a tared aluminum weighing dish, and place the dish overnight in a drying oven at 105°C. Weigh the oven-dried soil, and compute the soil water content, using the following formula (Gardner, 1965): 9/0 H,O = (g wet soil - g dry soil) x 1OO/(g dry soil - g dish)

Allocate the remainder of the soil sample as follows: 10 g for microbial diversity analysis, 6 g for dehydrogenase assay, and 1 g each for acid phosphatase, alkaline phosphatase, and arylsulfatase assays. Use


17

the remainder of the soil for the assay of other enzymes or other parameters (e.g., ATP content), as desired. Ideally, all assays should be conducted immediately after removal of the vials from the master jars. However, there is seldom sufficient technical assistance available to do this. Consequently, the investigator should determine which assays need to be conducted immediately and which can be conducted the following day. In some of the studies reported below, the assays for the phosphatases and sulfatases were conducted on the day of sampling, and the assays for dehydrogenases and microbial diversity, as well as for the other parameters, were conducted the following day after storage of the soil samples at 4"C. For analysis of microbial diversity, add 10 g of soil to 95 or 100 ml of sterile tap water in a 130-ml French square bottle, and shake on a rotary shaker at 160 rpm for 40 minutes (glass beads, 2 or 3 mm, can be added to enhance dispersion of the soil). Alternatively, use 250-ml Erlenmeyer flasks, and shake on a wrist-action shaker. Aseptically transfer 10 ml of the soil suspension to 90 ml of sterile tap water, and repeat until the desired series of dilutions is achieved. Immediately before each serial decade dilution, shake the dilution bottles vigorously for 15 to 20 seconds. Transfer 0.1ml of the desired dilutions to prepoured agar plates containing the appropriate selective medium, distribute the inoculum uniformly over the surface of the agar with a spreader (e.g., bent glass rod), invert, and ihcubate the plates in the dark at 24 ? 2°C for the prescribed period of time. Fungal plates are not inverted (Koch, 1981;Wollum, 1982).Count the number of colonies on plates containing 30 to 300 well-separated colonies after appropriate periods of incubation, and convert the numbers to CFU per gram soil, oven-dry equivalent, by multiplying the number of CFU by the dilution plated (remember that plating 0.1 ml constitutes another 10-fold dilution) and by 1 plus the percent soil water content (expressed as a decimal). For example, if 1 g of soil at the - 33-kPa water tension contains 0.25 g of water and 5.2 x lo8 CFU of total bacteria, the number of CFU/g oven-dry soil is 6.9 X lo8 [(l g - 0.75 g) X 100/0.75g = 33% water; (5.2 x lo8 CFU) x 1.33 = 6.9 x lo8 CFU/g oven-dry soil]. The battery of techniques described below for determining species diversity is not only reliable and reproducible, but it is also sensitive enough to detect even small changes in the diversity of species present in the soil microbiota (e.g., Bewley and Stotzky, 1983a,b,d; Stotzky and Goos, 1965,1966;Stotzky eta]., 1962,1966). Measure the soil pH with a pH meter on the 10-l dilution remaining

18

G . STOTZKY ET AL.

after making the l o - * dilution. To enhance sedimentation of clay particles, add 5 ml of a 0.5 M CaCl, solution (McLean, 1982). To determine both the contribution of added GEMs or the homologous parental strains to enzyme activities in soil and the effect of the indigenous soil microbiota on the survival of the GEMs and their homologous parents, comparative studies should be conducted in sterile soil. To conduct studies with sterile soil, place 50 g of soil, oven-dry equivalent, at the -33-kPa water tension into 100-cm3vials, plug the vials with cotton or glass wool, and weigh the vials. Autoclave the vials at 15 psi and 121°C for 15 min, allow to cool to room temperature, and autoclave again for 15 min. Aseptically add some soil from the autoclaved vials to nutrient broth, and check for the absence of turbidity after several days, to ensure that the soil was sterile. Alternatively, place crumbs of the autoclaved soil on plates of nutrient agar; no colonies should form if the soil is sterile. Place the sterile vials at room temperature (24 f ZOC) for 1 week in a humidified chamber, to allow for the dissipation of toxic compounds (Stotzky, 1973), and then weigh the vials to determine the amount of water lost. Adjust the soils to the -33-kPa water tension with sterile tap water containing the desired bacteria, prepared as described above. Mix the soil with a sterile applicator stick or spatula, and incubate the vials for 1 week at 24 f 2°C before conducting enzymatic or microbial analyses. Autoclave or otherwise sterilize all soils and soil dilutions before disposal. D. MICROBIAL ASSAYS 1. Maintenance Medium

Luria Agar Ingredient Tryptone Yeast extract NaCl Agar Distilled water

PH

Amount 10.0 g 5.0 g 5.0 g

15.0 g 1000 ml 7.0

Procedure. Use Luria agar (L-agar) for the storage of GEMs and their homologous parents. (For some bacteria, other media may be more appropriate.) Add the appropriate antibiotics, heavy metals, or other se-


19

lective agents to maintain the genes of interest. Transfer cultures regularly (e.g., every 2 to 4 weeks) to fresh medium. When L-agar is used for the recovery of the GEMs or parents from soil, add 100 to 250 mg/liter cycloheximide, before autoclaving (15 min at 15 psi and 121"C), to control fungal growth (the optimum concentration of cycloheximide should be determined for the soils being used). Add appropriate filtersterilized (0.2-pm filter) selective agents (see Table 111) for the selective recovery of the GEMs or parental strains after autoclaving and cooling the medium to about 50°C in a water bath. Aseptically dispense the medium into sterile petri plates, allow to solidify, invert, and dry overnight at 24 & 2°C. Replace the plates into their plastic storage sleeves, and store until use at 4°C in the dark, to reduce photodegradation of light-sensitive antibiotics. Count the plates after incubation at 24 & 2" C for 3 to 7 days. Discard unused plates after 5 days. 2. Bacteria (Total and Spore Formers)

Soil Extract Agar Ingredient

Amount

KZHPO. Dextrose Agar Soil extract Tap water

0.2 g 1.0 g 15.0 g 100 ml 900 ml

Procedure. Prepare soil extract by adding 500 ml of tap water to 500 g of soil in a 2-liter Erlenmeyer flask. Stopper the flask with a cotton plug or cover with aluminum foil, mix the suspension by swirling the flask vigorously, and autoclave for 60 min at 15 psi and 121°C. Allow the flask to cool, add 0.5 g of CaCO,, and vacuum-filter the suspension through Whatman No. 2 filter paper and a layer of diatomaceous earth or glass wool in a Buchner funnel. Adjust the volume of the filtrate to 500 ml with tap water and the pH to between 6.8 to 7.0 with 0.5 N HC1 or NaOH. Transfer aliquots (100 ml) of the filtrate to 130-ml French square bottles, autoclave at 121°C for 15 min, cap, and store at 4°C (James, 1958; Stotzky et al., 1962; Wollum, 1982). To prepare the medium, combine the ingredients, dissolve the agar on a hot plate with stirring, add cycloheximide (100-250 mg/liter) to control fungal growth, and autoclave at 121°C for 15 min. Store the poured plates at 4°C in the dark. When control of fungal growth is important, use the plates within 7 days for effectiveness of the cyclohexi-

20

G.STOTZKY ET AL.

mide (i.e., for total bacteria, use plates within 7 days; for spore-forming bacteria, where fungal growth is not a problem, use plates within 14 days). Enumerate total bacteria and spore-forming bacteria after incubation at 24 2°C for 5 to 10 days. For the enumeration of spore-forming bacteria, heat 10 ml of the appropriate serial dilutions of the soil at 80°C for 10 min. Place thermometers in several test tubes containing 10 ml of tap water, place these tubes and the tubes containing the soil dilutions in boiling water, and begin timing when the thermometers read 80°C. After exactly 10 min at 80°C, remove the tubes and immediately cool under running cold tap water.

*

3. Gram-Negative Bacteria

MacConkey Agar Ingredient

Amount

MacConkey agar Distilled water

45.0 g 1000 ml

Procedure. MacConkey agar is commercially available from Difco Laboratories. Dissolve the agar on a hot plate, with stirring, in distilled water, and autoclave at 121°C for 15 min. Allow the medium to cool to about 5O0C, and add appropriate antibiotics or other selective agents for the selective recovery of bacteria containing the novel genes of interest. When recovery is from nonsterile soil, add 100 to 250 mg/liter cycloheximide to inhibit fungi. Store the plates at 4°C in the dark, and use within 7 days. Examine the plates after 1 and 2 days. The colonies of lactose-positive strains of Escherichia coli or of other gram-negative species are dark red. 4. Pseudomonas putida

TNA Agar Ingredient

Amount

Tryptone Yeast extract Dextrose NaCl

5.0 g 2.5 g 1.0 g 8.5 g 20.0 g 1000 ml

Agar

Distilled water


21

Procedure. Combine the ingredients, dissolve the agar on a hot plate with stirring, and autoclave at 121°C for 15 min. Allow the medium to cool to about 50"C, and add appropriate antibiotics or other selective agents (Olsen and Shipley, 1973).When the recovery is from nonsterile soil, add 100 to 250 mg/liter cycloheximide to control fungal growth. Store the plates at 4°C in the dark, and use within 7 days. Count the plates after incubation at 24 f 2°C for 1to 2 days. 5. Fungal Propagules

Rose Bengal-Streptomycin Agar Ingredient

Amount

Dextrose Peptone KHZPO, MgSO4.7HzO Rose Bengal Agar Streptomycin Tap water

10.0 g 5.0g 1.0g 0.5 g 0.033g

20.0 g 2.4 ml 1000.0 ml

Procedure. Rose Bengal-streptomycin agar is used for the enumeration of fungal propagules in soil (Martin, 1950; Parkinson, 1982). The streptomycin and the acidic pH impair the growth of bacteria, and the Rose Bengal retards the spread of fungi. Prepare a sterile solution of streptomycin (1.25 mg/ml) by filtration (0.2-pm filter), store at 4"C, and add to autoclaved medium (15 min at 121°C) after cooling to about 50OC. The plates can be used up to 14 days after preparation. Count the plates after incubation (not inverted) at 24 2°C for 3 to 5 days.

*

Cellulose Utilizers Cellulose Agar

6.

Ingredient

Amount

KzmO4 NHINO, MgSO4.7HZO Microcrystallinecellulose (20 pm) Agar Soil extract Tap water

0.50 g 0.15 g 0.25 g 1.25 g 20.00 g 100 ml 900 ml

22

G. STOTZKY ET AL.

Procedure. Mix the KH2P04,NH4N03,MgS04.7H,0, and microcrystalline cellulose in 900 ml of tap water. Use 20-pm microcrystalline cellulose (Sigmacell Type 20 S-3504, Sigma Chemical Company, St. Louis, MO) instead of absorbent cotton treated with sulfuric acid (Harmsen, 1946). Add the soil extract (see Soil Extract Agar for preparation), cycloheximide (100-250 mg/liter), and agar, autoclave for 15 min at lZl"C, and store the plates at 4°C in the dark until use. Count the plates after incubation at 24 2°C for 7 to 14 days. Organisms that utilize cellulose form colonies surrounded by a zone of clearing of the agar, which, however, remains slightly clouded.

*

7. Chitin Utilizers

Chitin Agar Ingredient

KZHPO, KH2P0, MgSO,.7HZO FeS0,.7H20 ZnSO, MnCl Colloidal chitin (filter cake] Agar Tap water

Amount 0.700 g 0.300 g 0.500 g 0.010 g 0.001 g 0.001 g 4.000 g 20.000 g 1000 ml

Procedure. Grind 40 g of practical grade poly-N-acetylglucosamine, derived from crab shells (Chitin C-3387, Sigma Chemical Company), in a blender, and digest by stirring for 60 min in 400 ml of concentrated HC1. Precipitate the chitin as a colloidal suspension by adding the HCl digest slowly to 2000 ml of tap water chilled to 5 to 10°C. Collect the suspension by filtration, with suction, on Whatman No. 2 filter paper in a Biichner funnel, resuspend in 5000 ml of tap water, and refilter. Alternatively, collect the suspension by centrifugation at 5000 to 6000 g for 5 min. Repeat the washing procedure until the pH of the suspension is 3.5 (3 to 5 washings). Approximately 85% of the chitin should be recovered. Determine the water content of the chitin filter cake by drying a sample at 105°C. Autoclave and store the moist filter cake in a glass or ceramic jar covered with Parafilm and aluminum foil at 4°C until use (Hsu and Lockwood, 1975; Williams and Wellington, 1982). For use, add sufficient tap water to resuspend the chitin, and blend the suspension at high speed for 10 min. Dissolve the inorganic salts in 800 ml of tap water, add the chitin


23

suspension, cycloheximide (100-250 mglliter), and agar, bring to 1000 ml with tap water, adjust the medium to pH 8.0 with 5 N NaOH, autoclave for 15 min at 121"C, and store the plates at 4°C in the dark until use. Count the plates after incubation at 24 2°C for 7 to 14 days. Organisms that utilize chitin form colonies surrounded by a zone of clearing of the agar, which, however, remains slightly clouded. Most of the chitinoclastic colonies will consist of actinomycetes. The presence of actinomycetes can be confirmed by microscopic examination and by the cohesiveness of the colonies when poked with an inoculation needle. Colonies of actinomycetes will remain firm, whereas colonies of other bacteria will be disrupted and will spread.

*

Denitrifying Organisms Denitrifying organisms are evaluated by their ability to reduce NO,to a gaseous form of nitrogen (denitrifiers) or to reduce NO,- to NO,(nitrate reducers). 8.

Nitrate Broth Ingredient

Amount

Beef extract Peptone

3.0 g 5.0 g 1.00 g 1000 ml

mo3 Tap water

Bray's Nitrate-Nitrite Powder (Dry Mixture)

0

Ingredient

Amount (g)

MnS04*H20a Zinc dust" Sulfanilic acid a-Naphthylamine BaSO, Citric acid

10 2 4 2 100 75

Reducing agents are selectively added to Bray's powder to detect the presence of NO3-.

Reagents Glacial acetic acid (ACS reagent grade)

24

G. STOTZKY ET AL.

Procedure. Dissolve the beef extract and peptone (or use 8 g of Bactonutrient medium) and KNO, in 1000 ml of tap water. Dispense 5 ml of the solution into screw-capped test tubes, and autoclave for 15 min at 121°C. Alternatively, sterilize the nitrate broth in batch, and dispense 0.9 ml aseptically into sterile 1.8-ml microcentrifuge (e.g., Eppendorf) tubes. Add 0.1 ml each of at least four serial dilutions of the soil [e.g., 10-3, 10-4, 10-5, 10-6) to five replicate tubes containing either 5 or 0.9 ml of nitrate broth. Prepare Bray's nitrate-nitrite powder by mixing the MnS04-H20,zinc dust, sulfanilic acid, and a-naphthylamine and grinding thoroughly with a mortar and pestle, and then mix in the BaSO, and citric acid. Wear gloves and a dust mask or respirator, and prepare the powder in a vented fume hood (a-naphthylamine is carcinogenic, and Zn dust has a potential for explosion). N-(1-Naphthy1)ethylenediamine can be used instead of a-naphthylamine, but, as it is not known whether this compound is also carcinogenic, care should also be exercised in its use (Schmidt and Belser, 1982). Store the powder in a bottle covered with black tape or aluminum foil to exclude light. To distinguish between denitrifiers and nitrate reducers, exclude MnSO, and Zn dust from the powder when enumerating denitrifiers (Focht and Joseph, 1973). After incubation, at 24 & 2°C for 14 days, test for denitrification with Bray's nitrate-nitrite powder. Decant the test tubes (autoclave the decanted material before disposal; this is not necessary when using microcentrifuge tubes) to approximately a l-ml volume, and add 0.1 ml of glacial acetic acid and small amounts (-10 mg) of Bray's powder (without the reductants MnSO, and Zn dust). The formation of a red color indicates the presence of nitrate reducers (i.e., NO,- has been reduced to NO,-, which has combined with a-naphthylamine and sulfanilic acid). The absence of a red color indicates one of two conditions: NO,has been reduced completely to some form of gaseous nitrogen (e.g., NO, NzO, N,) by denitrifiers; or NO3- was not reduced. The addition of Zn dust (-10 mg) resolves this question: the absence of a red color after addition of Zn dust indicates that denitrification to gaseous nitrogen occurred, whereas the presence of a red color indicates that NO,- was not biologically reduced but was reduced by the Zn dust to NO,-.Calculate the numbers of denitrifying and nitrate-reducing organisms from most probable number (MF")tables (e.g., Cochran, 1950). 9. Nitrifying Organisms

Nitrifying organisms are evaluated by their ability to oxidize NH,' to NO,- (ammonium oxidizers) or NO,- to NO,- (nitrite oxidizers).


Ammonium-Oxidizer Broth

Ingredient

Amount (g) per 100 ml stock solution

Amount (ml) stock

5.00 10.0 (NWzSO, CaClz~2H,00 1.34 1.o MgS0,.7H,0a 4.00 1.0 Chelated iron 1.0 FeS0,.7H20 0.25 Na, EDTA 0.33 Trace elements 1.0 NaMoO,.ZH,O 0.01 MnCl, 0.02 CoCl,~6HzO 0.0002 ZnS04-7H,0 0.01 CuSO,*5H,O 0.002 Bromthymol blue 0.04 5.0 Distilled water 73.5 Use 5% NaOH to adjust the medium to pH 7.0 to 7.4 before sterilization a Materials are combined, autoclaved separately, and added aseptically to an autoclaved solution of the other ingredients.

Nitrite-Oxidizer Broth

Ingredient KNOZ CaCl,.2H,0° MgS0,.7H200 K2HPO4

Amount (8) per 100ml stock solution 0.85 1.34 4.00 3.48 2.72

Amount (ml) stock 1.0 1.0 5.0 4.0

1.0 KHZPO, 1.0 Chelated iron FeS0,.7Hz0 0.25 Na, EDTA 0.33 Trace elements 1.0 NaMoO,.ZH,O 0.01 MnCl, 0.02 CoC1,.6H2O 0.0002 ZnS0,.7H20 0.01 CuS0,.5H,O 0.002 Distilled water 986.0 Use 5% NaOH to adjust the medium to pH 7.2 to 7.5 before sterilization 0 Materials are combined, autoclaved separately, and added aseptically to an autoclaved solution of the other ingredients.

25

26

G. STOTZKY ET AL.

Reagents Glacial acetic acid (ACS reagent grade) Bray's nitrate-nitrite powder (dry mixture) (described under Denitrifying Organisms) Procedure. Dissolve constituents individually in 100 ml of distilled water, add appropriate aliquots of the stock solutions to a 1000-ml volumetric flask, and bring the volume to 1000 ml with distilled water. Autoclave the solutions in an Erlenmeyer flask for 15 min at 121°C, and mix the solutions after cooling (Schmidt and Belser, 1982). Aseptically add 0.24 ml of the broth to each well of a sterile microtiter plate (8 x 12 wells) (use a sterile Cornwall repeating syringe dispenser or a multichannel micropipetter). Add 0.06 ml each of at least four serial dilutions of the soil (e.g., 10-,, 10- 4 , 1 0 -5,10 -6) to five replicate wells containing the appropriate medium with sterile 1-ml pipettes, Stack the inoculated microtiter plates on top of each other, cover the top plate with a sterile microtiter plate lid, wrap the stacks in aluminum foil or Parafilm, to minimize loss of moisture, place the stacks in Styrofoam boxes (e.g., inner dimensions 20.6 x 15.5 x 16.5 cm; only -60% of the volume is occupied by the plates, to ensure adequate OJ, and incubate for 42 days at 24 ? 2°C. Determine the numbers of ammonium oxidizers by adding 0.03 ml of glacial acetic acid and a small amount (-10 mg) of Bray's nitrate-nitrite powder, containing MnSO, and Zn dust, to each well of the plate containing ammonium-oxidizer broth. The formation of a red color in the wells indicates the oxidation of NH,' to NO,- or NO,-. Determine the numbers of nitrite oxidizers by adding similar amounts of glacial acetic acid and Bray's powder, without MnSO, and Zn dust, to each well of nitrite-oxidizer broth. The absence of a red color is presumptive that NO,- has been oxidized to NO3-. Add Zn dust to the wells that show no color. If they now turn red (indicative of the chemical reduction of NO3- to NOz-), they are considered to be positive for the oxidation of NO,-. Subtract the numbers of nitrite oxidizers from the numbers of apparent ammonium oxidizers (which include both ammonium and nitrite oxidizers) to estimate the numbers of microbes that oxidized ammonium to nitrite. Calculate the numbers of ammonium-oxidizing and nitrite-oxidizing organisms from MPN tables (e.g., Cochran, 1950). 10. Protozoa

Correlations between the survival of the introduced bacteria, with and without the novel genes, and fluctuations in the numbers of protozoa should provide some insight into the effect of predation on the survival of the GEMS or the homologous parents in soil.


27

Protozoa Water Agar Ingredient

Amount

NaCl Agar Tap water

5.0 g 10.0 g

1000.0ml

Procedure. Dissolve the NaCl in 1000 ml of tap water, add and dissolve the agar, and autoclave for 15 min at 121°C (Singh, 1946). Aseptically dispense the medium into sterile petri plates, allow to solidify, invert, and dry overnight at 24 k 2°C. Core the plates with the open end of a test tube (25 mm OD) that has been flamed with ethanol, so that islands of agar, 25 mm in diameter, are physically separated from the remaining agar. Spread over each island a loopful of a suspension of mixed bacteria to serve as a food source for the protozoa. (Pick colonies from soil extract agar plates, and mix with a loop in 1 to 3 ml of tap water until a viscous slurry is formed.) Add 0.1 ml each of at least four serial soil dilutions (e.g., 10-7 to the center of lo-, five replicate islands. Incubate the plates, without inverting, in the dark for 5 days at 24 & 2°C. Examine each island under low-power magnification [ x 100) for the presence of protozoa, both vegetative and encysted forms. Clearing of the inoculated biomass is presumptive for the presence of protozoa, and visual observation of protozoa (flagellates, ciliates, and amoebas) is used to confirm their presence. Calculate the numbers of protozoa from MPN tables (e.g., Cochran, 1950). The numbers of protozoa in soil can also be estimated by the MPN method using a liquid medium (Allen, 1951). Prepare hay infusion broth by boiling 50 g of dry hay in 3.5 liters of tap water for 2 hours: cool overnight, filter (Whatman No. 2), dilute to 5 liters with tap water, autoclave for 15 min at 121OC. and store at 4°C. Aseptically add either 0.27 ml of the broth to each well of a microtiter plate or 9 ml to screwcapped test tubes, and add 0.03 or 1 ml, respectively, of at least four serial soil dilutions to five replicate wells or tubes. Incubate the tubes on a slant, and do not screw the caps on tightly, to enhance aeration of the medium. Incubate for at least 7 days at 24 & 2°C. If time permits, enumerate again after 14 and 2 1 days, and attempt to enumerate each group of protozoa (e.g., flagellates, ciliates, amoebas) separately. Examine the wells directly or 0.1-ml aliquots from the test tubes under lowpower magnification ( x 100) for the presence of protozoa. Calculate the numbers of protozoa from MPN tables (e.g., Cochran, 1950), as in the plate counts. The counts obtained by the solid and liquid media methods are similar, as indicated in Table 11.

28

G. STOTZKY ET AL. TABLE I1

COMPARISON OF NUMBERS OF PROTOZOA ENUMEMTED ON SOLID OR IN

GEM or homologous host added to soil Control (Hz0) E. coli W3110 E . coli W3110(R702)

Number of protozoa x

LIQUIDMEDIAO

(solid/liquid)

Day 1

Day 7

Day 14

2.1112.46 2.3212.44

2.0112.22 2.09/2.02

2.4212.51 2.4412.35

1.8411.97

2.1412.23

2.6612.59

Osee text for details.

11. Physiological Groups

The effects of GEMS on the diversity of the soil microbiota can also be estimated by measuring changes in physiological (nutritional) groups as an indicator of changes in bacterial populations and community structure. A dynamic equilibrium exists between various physiological groups of bacteria in soil. This equilibrium depends, in part, on the nutrients available. The availability of nutrients may, in turn, be affected by an introduced GEM, which, because of its novel genetic material, could compete effectively with the indigenous bacteria for specific nutrients and/or could produce substances that disrupt nutrient uptake and use. Changes in the nutritional status of soil bacteria can be determined by the use of modifications of the procedures developed by Lochhead and Chase (1943).The growth of bacteria isolated from soil inoculated with the GEM or the homologous parent or not inoculated is compared on or in media of varying complexity. Inasmuch as just the introduction of a quantity of bacteria into soil may impact the nutritional balance, changes in the nutritional status of indigenous bacteria in soil inoculated with the homologous parental strain must be evaluated as a control. Basal Medium Ingredient Dextrose KzHPO, KNO, MgSO,*7HzO CaCl, FeC1,.6Hz0 Agar Distilled water

Amount 1.00 g 1.00g 0.50g

Amount (g) per 4 liters 8.00 4.00

0.20 g

0.10g 0.01 g 15.00 g 1000 ml

0.80 0.08

120.00


29

Prepare a 0.1 g/ml solution of dextrose and a 0.2 g/ml solution of MgS04.7H,0 in distilled water. Dissolve the K,HP04 and KNO, in 4 liters of distilled water (solution A]. Dissolve the CaCl,, NaCl, and FeC1,.6H2O (and agar, if plates are desired) in 4 liters of distilled water (solution B). Autoclave all solutions for 30 min at 121OC. Store component solutions at 4' C. Mix 10 ml of the dextrose solution, 494.5 ml of solution A, 1 ml of the MgS04.7H,0 solution, and 494.5 ml of solution B. Adjust the final 1000-ml mixture to pH 6.8 with 0.1 N HCl or 0.1 N NaOH, and sterilize by passage through a 0.2-pm filter. Basal plus Amino Acids Medium Ingredient

Amount

Vitamin assay casamino acids Basal medium

5.0 g 1000 ml

Dissolve the vitamin assay casamino acids (Difco) in 100 ml of solution A of the basal medium, and sterilize by passage through a 0.2-pm filter. Add the casamino acid solution to 394.5 ml of sterile solution A, mix, and aseptically add the other components of the basal medium, as described above. Basal plus Growth Factors Medium Ingredient Cysteine Thiamine Biotin Pyridoxine Pantothenic acid Nicotinic acid Riboflavin Inositol Basal medium

Amount 0.0500000 g

0.0001000g 0.0000001g 0.0002000 g 0.0001000g 0.0001000g 0.0002000 g 0.0500000 g 1000 ml

Dissolve the individual growth factors in 100 ml of solution A of the basal medium, and sterilize by passage through a 0.2-pm filter. Add the growth factor solution to 394.5 ml of sterile solution A, mix, and aseptically add the other components of the basal medium, as described above.

30

G. STOTZKY ET AL.

Basal plus Amino Acids and Growth Factors Medium Ingredient

Amount ~~~~

Vitamin assay casamino acids Cysteine Thiamine Biotin Pyridoxine Pantothenic acid Nicotinic acid Riboflavin Inositol Basal medium

5.0000000 g 0.0500000 g 0.0001000 g 0.0000001 g 0.0002000 g 0.0001000 g 0.0001000 g 0.0002000 g 0.0500000 g

1000 ml

Dissolve the vitamin assay casamino acids and growth factors in 100 ml of solution A of the basal medium, and sterilize by passage

through a 0.2-pm filter. Add the casamino acid-growth factor solution to 394.5 ml of sterile solution A, mix, and aseptically add the other components of the basal medium, as described above. Yeast Extract Medium Ingredient

Amount

Yeast extract Basal medium

1000 ml

1g

Dissolve the yeast extract in 100 ml of solution A of the basal medium, and sterilize by passage through a 0.2-pm filter. Add the yeast extract solution to 394.5 ml of sterile solution A, mix, and aseptically add the other components of the basal medium, as described above. Soil Extract Medium Ingredient

Amount (ml)

Soil extract Basal medium

250 750

Prepare the basal medium as described above. Add 250 ml of filtersterilized (0.2-pm filter) soil extract (see Soil Extract Agar for preparation) to 750 ml of sterile basal medium and mix.


31

Yeast Extract and Soil Extract Medium Ingredient

Amount (ml]

Soil extract Yeast extract medium

250 750

Prepare the yeast extract medium as described above. Add 250 ml of filter-sterilized (0.2-pm filter) soil extract to 750 ml of sterile yeast extract medium and mix. Yeast Extract and Vitamin B,, Medium Ingredient Vitamin Blz Yeast extract Basal medium

Amount 0.000002 g 1.oooooo g 1000 ml

Dissolve the vitamin BIZ and yeast extract in 100 ml of solution A of the basal medium, and sterilize by passage through a 0.2-pm filter. Add the vitamin B,,-yeast extract solution to 394.5 ml of sterile solution A, mix, and aseptically add the other components of the basal medium, as described above. Procedure for Assaying Physiological Groups. Prepare the following agar (1.5 or 2% agar) media: basal and basal plus amino acids, growth factors, amino acids and growth factors, yeast extract, soil extract, yeast extract and soil extract, and yeast extract and vitamin BIZ. Dry the media overnight at 24 & 2°C. Transfer at least 50 representative and well-separated colonies with sterile toothpicks from countable (30 to 300 colonies) soil extract agar plates to individual wells in microtiter plates containing 0.3 ml of sterile saline (0.85% NaC1). Inoculate at least two wells with each colony, twirling the toothpick 10 times in the well. Use a new toothpick for each colony. Autoclave the toothpicks before disposal. Inoculate all isolates in a microtiter plate simultaneously into the appropriate agar media with a needle replicator (Stotzky, 1965b). Inoculate two agar plates of each medium for each charging of the replicator: place the replicator gently on the agar in the first plate, remove, and then push the needles through the agar in the second plate. Sterilize the replicator between inoculation of the different media by

32

G.STOTZKY ET AL.

dipping in ethanol and flaming. Allow the needles to cool before charging. Incubate the agar plates for 5 days at 24 ? 2°C. Evaluate each plate for (1)the presence of colonies and (2) colony size on a scale of one to four (four being the largest). After inoculation of all the media plates, seal the microtiter plates containing the original isolates with Parafilm, and store at 4°C for later reference. Alternatively, use liquid media of the same composition as the agar media, Transfer 0.3 ml of the appropriate liquid medium to duplicate wells of a sterile microtiter plate. Inoculate the medium-containing microtiter plates with the needle replicator, as above. Use a microtiter plate reader to evaluate growth on the basis of turbidity. Compare the growth of microorganisms isolated from uninoculated soil and from the same soil inoculated with either the parental or GEM strains. 12. Antibiotic-Resistant Phenotypes

Profiles of antibiotic-resistant phenotypes can also be used as another indicator of the effects of GEMS on microbial communities in soil. These profiles in soil can vary, depending on the time of sampling and on the physicochemical and biological characteristics of the soil (Stotzky, 1972).Perturbation of any of these characteristics, as might occur after the introduction of a GEM, could alter the profile of antibioticresistant bacteria. Changes in the resistance to antibiotics of the soil microbiota, in the presence of a GEM, may indicate (1)the expression of the novel genes by the GEM; (2)the acquisition and expression of antibiotic-resistance genes from the GEM by the indigenous microbiota; (3) the expression of unanticipated pleiotropic effects resulting from the acquisition of novel genetic elements; and/or (4)changes in soil conditions and microbial populations as the result of the GEM or its metabolic products (e.g., Kozyrovskaya et a]., 1984;Stotzky, 1989). Shifts in the antibiotic-resistant phenotypes of the soil microbiota can be determined by methods similar to those used for evaluating changes in the physiological groups of the microbiota. The growth of bacteria isolated from uninoculated soil on or in media containing different antibiotics at various concentrations is compared, over time, with the growth of bacteria isolated from the same soil that has been inoculated with the GEM or the homologous parental strain. The antibioticresistant phenotypes to be monitored depend on (1)the genotype of the GEM; (2)the present and historic land-use practices followed at the location where the soil was collected; and (3)the profile of antibioticresistant phenotypes found in the unamended and uninoculated soil in preliminary studies. Each antibiotic should be tested at relatively high, medium, and low concentrations.


33

Antibiotics. Various antibiotics can be used for evaluating antibioticresistant phenotypes in soil, and no attempt is made here to give specific formulations. Some guidelines are presented in Table 111. In general, all antibiotics should be filter-sterilized (0.2-pm filter) before use. Storage time and temperature are dependent on the antibiotic (refer to the manufacturer's specifications or the most recent Merck Index). All media should be cooled to approximately 50°C in a water bath before adding the appropriate antibiotic(s). The following antibiotics are only illustrative of those used in these studies. Procedure. Prepare soil extract agar plates containing the following final concentrations of each filter-sterilized antibiotic: 50, 100, and 150 pg/ml carbenicillin; 10, 30, and 90 pg/ml chloramphenicol; 10,30, and 90 ,ug/ml kanamycin; 1, 10, and 100 pg/ml streptomycin; and 10, 30, and 90 pg/ml tetracycline. [The antibiotic concentrations chosen were based on those recommended by Bauer et al. (1966) and those used by Kozyrovskaya et al. (1984).] Transfer at least 50 representative and well-separated colonies with sterile toothpicks from countable (30 to 300 colonies) soil extract agar plates to individual wells in microtiter plates containing 0.3 ml of sterile saline (0.85% NaCl). Inoculate at least two wells with each colony, twirling the toothpick 10 times in the well. Use a new toothpick for each colony. Autoclave the toothpicks before disposal. Inoculate all the isolates in a microtiter plate simultaneously into the appropriate agar media with a needle replicator (Stotzky, 1965b). Ideally, use the same microtiter plates containing the soil isolates that were evaluated for physiological groups. Inoculate two agar plates of each medium for each charging of the replicator: place the replicator gently on the agar in the first plate, remove, and then push the needles through the agar in the second plate. Sterilize the replicator between inoculation of the different antibioticcontaining media by dipping in ethanol and flaming. Allow the needles to cool before charging. Incubate the agar plates for 5 days at 24 & 2°C. Score each plate for the presence of colonies. Compare the resultant profiles over time. Seal the microtiter plates containing the original isolates with Parafilm, and store at 4OC for later reference. Alternatively, use liquid media containing the same concentrations of antibiotics. Transfer 0.3 ml of the appropriate liquid medium to duplicate wells of a sterile microtiter plate. Inoculate the mediumcontaining microtiter plates with the needle replicator, as above. Use a microtiter plate reader to evaluate growth on the basis of turbidity. Compare the sensitivity to each antibiotic of microorganisms isolated

TABLE III ANTIMICROBIAL AGENTS COMMONLY ADDEDTO MEDIAFOR SELECTIVE RECOVERYOF GENETICALLY ENGINEERDMICROORGANISMS FROM Son. AND FOR EVALUATING ANTIBIOTIC-RESISTANT PHENOTYPES ISOLATED FROM SOIL Concentrated st&

Concentration in medium Amount (ml) to add to500ml

Amount (ml)to add tOl00ml

Amount (ml) to add t o l d

0.200 0.100 0.500

4.00

0.80 0.20 1.00

0.010 0.050

0.025 0.025 0.015 0.025 0.025 0.050

1.00 0.50 0.75

Stability

Li& ABent

Abbreviation sensitivity

Ampidin l3lbnicillin CephalOthin

JQ cb

cp

-

Chloramphenicol

cm

-

Chlortetracycline

ct

Cyclohsximide

CY

Gwtamicin Kanamycin

Gn Kn

Mercuric chloride Nafcillin Nalidixic acid

Nf Nx

w

Rifampicin Streptomycin Sulfemxisole Tetracycline Trimethoprim

Rif

Thiostrepton Tobramycin

TI

Sm sx Tc TP

Tb

+ + -

+

Diluent Water Water 0.1 M phosphate,

PH 6 Ethanol 50% ( d v )ethanol Water Watar Water Water Water 0.1 M NaOH

-

+

Methanol Water 0.1 M NaOH 50% (v/v) ethanol

-

15% 0.06

-

HCI qs water Dimethyl sulfoxide Water

M

Concentration

at 4°C

Concentration

(ms/ml)

(days)

(ms/ml)

25.0 50.0 12.5

7 14 5

25.0 10.0 12.5 12.5 25.0

30 5 5 30 30 30 5 30

40.0

1.0 16.0 16.0 25.0 12.5 25.0

30 5

30

2.0

15 5 30

50.0 5.0

30

10.0

-

0.040

0.002 0.032

1.00 5.00

1.00 1.00 1.00 0.50 1.00

0.20

0.10 0.15 0.20 0.20 0.20

0.10

1.00

0.20 0.20

0.500 0.100 0.025 0.200 0.015 0.050

15.60 2.00 1.00 4.00 0.75 12.50

3.12 0.40 0.20 0.80 0.15 2.50

0.050 0.010

0.50 0.10

0.02

0.10

0.040

0.010 0.005 0.008

0.010 0.010 0.010 0.005 0.010 0.010 0.156 0.020 0.010 0.040 0.008 0.125

0.005 0.001

RELEASE OF GENETICALLY ENGINEERD MICROORGANISMS

35

from uninoculated soil and from the same soil inoculated with either the parental or GEM strains. If consistent significant differences are apparent between isolates from uninoculated soil and the soil inoculated with the GEM or the homologous parent, the antibiotic screen should be enlarged by the use of Sensititre multiple-antibiotic minimum inhibitory concentration (MIC) plates (GIBCO) or comparable commercial systems. 13. Biochemical Phenotypes

As another approximation of whether the introduction of novel genes results in unrelated and unanticipated pleiotropic effects (e.g., Kozyrovskaya et al., 1984;Stotzky, 1989),isolated soil bacteria can be subjected to simple biochemical screens. The initial screen for detecting altered biochemical characteristics consists of microtiter plates containing a liquid basal medium augmented with different carbohydrates (e.g., glucose, lactose, maltose, sucrose, mannitol, salicin, rhamnose, dulcitol, sorbitol) and phenol red. Using the same microtiter plates containing the soil isolates that were evaluated for physiological groups and antibiotic-resistant phenotypes, inoculate each isolate with the needle replicator into four wells of each carbohydrate medium. Layer two wells of each medium with mineral oil to provide anaerobic conditions. Alternatively, incubate inoculated microtiter plates containing two wells of each medium under aerobic and anaerobic conditions. The ability of the isolates to utilize and/or ferment the carbohydrates is determined by change in the color of the phenol red indicator. If consistent significant differences in utilization and fermentation patterns are observed between isolates from the uninoculated soil and from the soil inoculated with either the GEM or the homologous parent, the biochemical screens should be expanded (e.g., use additional carbohydrates and other biochemical characteristics commonly employed in the numerical taxonomy of environmental isolates). Commercial systems (e.g., API, Biolog) can be used for determining the utilizationlfermentation of various substrates. Moreover, all personnel involved in the studies should be alert for other unanticipated pleiotropic effects (ens.,increased production of capsular slime in soil isolates, filamentation, increased mutation frequencies, decreases in the utilization of common energy and carbon substrates) (e.g., Stotzky, 1989). 14. Fate of Novel Genes

The survival and fate of the introduced GEMS and of the novel genes (i.e., in the event of transfer to indigenous soil bacteria) should be evaluated in the subsamples of soil from the master jars (e.g., by phe-

36

G.STOTZKY ET AL.

notypic characterization on selective media, DNA fingerprinting before and after restriction by various endonucleases, DNA probes) (Devanas and Stotzky, 1986;Devanas et al.,1986;Jain et al., 1988;Kado and Liu, 1981;Krasovsky and Stotzky, 1987; Sayler et a]., 1991; Short et al., 1991;Stotzky, 1989;Stotzky et al., 1990;Zeph and Stotzky, 1989;Zeph et al., 1988,1991),to establish whether changes in microbial populations and processes are related to the presence of the GEMs or the novel genes. To determine whether residence in soil results in any changes in the antibiotic-resistance patterns or biochemical characteristics of the GEMs, colonies developing on media selective for the GEMs or their novel genes should be replica plated, either after direct isolation from soil or after transfer from a less stressful isolation medium (e.g., Stotzky et al., 1990),to antibiotic-containing agars and inoculated into media containing different carbohydrates, as detailed above. Any isolates showing significant differences in antibiotic-resistance patterns and/or biochemical characteristics should be identified by classic biochemical and molecular techniques (e.g., gel electrophoresis, restriction enzyme patterns) (Ausubel et al., 1987;Maniatis et al., 1982).

E. SOILENZYMES Subsamples of soil from the master jars should be analyzed for the activity of selected enzymes, as soil enzyme activity has often been used as an index of soil metabolism, especially when soils are stressed (e.g., Babich and Stotzky, 1983, 1985). The enzymes that should be studied initially are acid and alkaline phosphatases, arylsulfatases, and dehydrogenases. The activities of the phosphatases can be used to estimate the mineralization of organic phosphates by measuring colorimetrically the release of p-nitrophenol from p-nitrophenyl phosphate; a buffer at pH 6.5 is used to assay the acid phosphatase activity, and a buffer at pH 11 is used to assay the alkaline phosphatase activity (Tabatabai and Bremner, 1969).The activity of arylsulfatases, which catalyze the hydrolysis of arylsulfate anions, can be used to estimate the mineralization of organic sulfur by measuring colorimetrically the release of p-nitrophenol from potassium p-nitrophenyl sulfate (Tabatabai and Bremner, 1970).Dehydrogenase activity, which is another measure of the biological oxidation of organic compounds, can be estimated by colorimetric measurement of the concentrations of triphenylformazan that result from the reduction of 2,3,5-triphenyltetrazoliumchloride (Johnen and Drew, 1977). If significant differences (e.g., p < 0.05) in the activities of these enzymes are consistently observed between control soils and soils inoculated with a GEM or the homologous parent,


37

evaluation of the activities of additional enzymes, for example, amylases (Cole, 1977), cellulases (Spalding, 1979), proteases (Caplan and Faley, 1980), and nucleases (to provide information on the possibility of transforming DNA persisting in soil) (Tabatabai, 1982), should be considered. The activities of these and other hydrolytic enzymes should be evaluated when appropriate substrates (e.g., starches, celluloses) are added to the soils. Various methodologies have been used to study enzyme activities in soils, but there is some controversy about which methods are best (e.g., Burns and Slater, 1982; Nannipieri et a]., 1990; Skujins, 1976). The procedures of Tabatabai and co-workers (e.g., Al-Khafaji and Tabatabai, 1979; Eivazi and Tabatabai, 1977; Frankenberger and Tabatabai, 1982; Fu and Tabatabai, 1991; Juma and Tabatabai, 1977; Tabatabai, 1982) appear to be the most accepted methods. The caveat of Malcolm (1983) that the concentration of substrate added to soil be sufficiently high (e.g., at least 10 times the K, value) to saturate the enzyme system being studied should be considered in the enzyme assays. 1. Phosphomonoesterases (Acid and Alkaline Phosphatases)

Reagents Toluene (ACS reagent grade) Modified universal buffer (MUB): Dissolve 12.1 g tris(hydroxymethyl)aminomethane, 11.6 g maleic acid, 14.0 g citric acid, and 6.3 g H,BO, in 488 ml of 1 N NaOH, dilute to 1 liter with distilled water, and store at 4°C. Before use, adjust the pH of 200 ml of the solution with 0.5 N HCl to pH 6.5 for the assay of acid phosphatase or with 0.5 N NaOH to pH 11.0 for the assay of alkaline phosphatase. Adjust the volumes of the pH-modified buffers to 1 liter with distilled water. p-Nitrophenyl phosphate (PNP) substrate: Dissolve 0.46 g disodium p-nitrophenyl phosphate hexahydrate (Sigma 104, Sigma Chemical Company) in 40 ml of the appropriate pH-adjusted, diluted MUB (pH 6.5 for acid phosphatase and pH 11.0 for alkaline phosphatase), and dilute to 50 ml with MUB of the same pH. Store the PNP substrate at 4°Cfor no longer than 10 days. CaC1, (0.5 M): Dissolve 73.5 g CaC1,-2H,O in 700 ml of distilled water, and dilute to 1 liter with distilled water. NaOH (0.5 N): Dissolve 20.0 g NaOH in 70 ml of distilled water, and dilute to 1 liter with distilled water. p-Nitrophenol standard: Dissolve 1.0 g p-nitrophenol in 700 ml of distilled water, and dilute to 1 liter with distilled water. Store the solution in a dark bottle at 4°C for no longer than 21 days.

38

G. STOTZKY ET AL.

Procedure. Add 1.0 g of soil, 0.2 ml of toluene, and 4.0 ml of the diluted MUB (pH 6.5 for acid phosphatase and pH 11.0 for alkaline phosphatase) to a 50-ml Erlenmeyer flask. Add 1ml of the PNP substrate, at the same pH as the MUB, to the flask, swirl to ensure adequate mixing, stopper the flask (Parafilm works well), and place in a water bath at 37°C for 60 min. Remove the flasks in the sequence in which they were placed in the bath, add 1ml of the CaCl, solution and 4 ml of the NaOH solution to each flask, swirl, and filter the suspension through a folded, 100-mm disk of Whatman No. 2 filter paper in a 65-mm (top diameter) short-stem funnel with a fluted bowl. Controls are analyzed in a similar manner, except that the PNP substrate is added after the CaCl, and NaOH solutions. Add 1 g of soil, 0.2 ml of toluene, and 4.0 ml of MUB at the appropriate pH to a 50-ml Erlenmeyer flask, as described above. Stopper the flask and place in a water bath at 37°C for 60 min, add 1ml of the CaCl,, 4 ml of the NaOH, and 1ml of the appropriate PNP solutions, swirl, and filter, as above. Measure the absorbance of the filtrate as soon as possible after preparation with a spectrophotometer at 400 nm, and convert absorbance units to concentrations from a standard curve of p-nitrophenol. Prepare the standard curve as follows: dilute the stock solution of p-nitrophenol 1:100 in a volumetric flask with distilled water, mix well, transfer 0, 1, 2, 3, 4, and 5 ml of the diluted solution to 25- or 50-ml Erlenmeyer flasks, and adjust the volume to 5 ml with distilled water. Add 1 ml of the CaC1, and 4 ml of the NaOH solutions to each flask and filter, as above, to yield 0,10, 20, 30, 40,and 50 pg/ml p-nitrophenol. Plot the absorbance of the standards against their concentration, and convert the absorbance of the samples to concentration with the standard curve. Zero the spectrophotometer with the controls for the samples and with the 0-pg p-nitrophenol standard solution for the standards. If the absorbance of a sample exceeds that of the standards, dilute the sample with distilled water. Include this dilution when calculation the concentration of p-nitrophenol. Convert the data to an oven-dry basis by multiplying the p-nitrophenol value by 1 plus the percent soil water content (expressed as a decimal). Record the data as micrograms p-nitrophenol per gram soil, oven-dry equivalent. 2. Arylsulfatases

Reagents Toluene (ACS reagent grade) Acetate buffer: Dissolve 68 g sodium acetate trihydrate in about 700 ml of distilled water. Add 1.70 ml of glacial acetic acid (99%), and dilute to 1liter with distilled water. Store the buffer at 4"C for no longer than 14 days.


39

p-Nitrophenyl sulfate (PNS) substrate: Dissolve 0.322 g potassium pnitrophenyl sulfate in 40 ml of acetate buffer, and dilute to 5 0 ml with acetate buffer. Store the substrate at 4°C for no longer than 10 days. CaC1, (0.5 M): Dissolve 73.5 g CaC1,.2H,O in 700 ml of distilled water, and dilute to 1liter with distilled water. NaOH (0.5 N): Dissolve 20.0 g NaOH in 70 ml of distilled water, and dilute to 1liter with distilled water. p-Nitrophenol standard: Dissolve 1.0 g p-nitrophenol in about 700 ml of distilled water, and dilute to 1liter with distilled water. Store the standard in a dark bottle at 4°C for no longer than 2 1 days. Procedure. Add 1 g of soil, 0.25 ml of toluene, and 4.0 ml of acetate buffer to a 50-ml Erlenmeyer flask. Add 1.0 ml of the PNS substrate to the flask, swirl to ensure adequate mixing, stopper the flask, and place in a water bath at 37°C for 6 0 min. Remove the flasks in the sequence in which they were placed in the bath, add 1 ml of the CaC1, and 4 ml of the NaOH solutions to each flask, swirl, and filter the suspension through a folded, 110-mm disk of Whatman No. 2 filter paper in a 65mm (top diameter) short-stem glass funnel with a fluted bowl. Controls are analyzed in a manner similar to the samples, except that the PNS substrate is added after the CaC1, and NaOH solutions. Add 1g of soil, 0.25 ml of toluene, and 4 ml of acetate buffer to a 50-ml Erlenmeyer flask. Stopper the flask and place in a water bath at 37°C for 60 min, add 1 ml of the CaCl,, 4 ml of the NaOH, and 1 ml of the PNS solutions, swirl, and filter, as above. Measure the absorbance of the filtrate as soon as possible after preparation with a spectrophotometer at 400 nm, and convert absorbance units to concentrations from a standard curve of p-nitrophenol. Prepare the standard curve as follows: dilute the stock solution of p-nitrophenol 1:100 in a volumetric flask with distilled water, mix well, transfer 0 , 1, 2, 3, 4, and 5 ml of the diluted solution to,25- or 50-ml Erlenmeyer flasks, and adjust the volume to 5 ml with distilled water. Add 1 ml of the CaCl, and 4 ml of the NaOH solutions to each flask and filter, as above, to yield 0,10, 20, 30, 40, and 50 pg/ml p-nitrophenol. Plot the absorbance of the standards against their concentration, and convert the absorbance of the samples to concentration with the standard curve. Zero the spectrophotometer with the control for the samples and with the 0-pg p-nitrophenol standard solution for the standards. If the absorbance of a sample exceeds that of the standards, dilute the sample with distilled water. Include this dilution when calculating the concentration of p-nitrophenol. Convert the data to an oven-dry basis by mul-

40

G.STOTZKY ET AL.

tiplying the p-nitrophenol values by 1 plus the percent soil water content (expressed as a decimal). Record the data as micrograms pnitrophenol per gram soil, oven-dry equivalent. 3. Dehydrogenases

Reagents CaCO, (powder, reagent grade) 3,5-Triphenyltetrazoliumchloride (TTC);3% (w/v) in water Methanol (ACS certified) Triphenylformazan (TPF):Dissolve 0.10 g TPF in 80 ml of methanol, and adjust to 100 ml with methanol. Prepare new standards for each assay, as TPF is light-sensitive. Procedure. Add 6 g of soil and 0.06 g of CaCO, to a screw-capped test tube (16 x 150 mm). Add 1.0 ml of the TTC solution and 2.5 ml of distilled water to the tube, cap, vortex, and place in a water bath at 37"C for 24 hours. Remove the tubes in the sequence in which they were placed in the bath, add 10 ml of methanol, mix for 1 min, and completely transfer the soil suspension from the tube with a series of methanol rinses to a vacuum-filtration unit, containing Whatman No. 2 filter paper in a 55-mm diameter Buchner funnel. Wash the, soil with aliquots of methanol until no more red color is removed from the soil. Combine the methanol rinses in a 100-ml volumetric flask, adjust the final volume to 100 ml with methanol, and mix by inversion (Tabatabai, 1982). Measure the absorbance with a spectrophotometer at a wavelength of 485 nm, using methanol as a blank, and convert the absorbance units to concentrations of TPF from a standard curve of TPF. Prepare the standard curve by diluting the TPF solution 1:10 with methanol and diluting 5,10,15, and 20 ml of this TPF solution to 100 ml with methanol in volumetric flasks to yield 5, 10, 15, and 20 pg/ml TPF. Zero the spectrophotometer with a methanol blank. Convert the data to an ovendry basis by multiplying the TPF values by 1plus the percent soil water content (expressed as a decimal). Record the data as micrograms TPF per gram soil, oven-dry equivalent. Some investigators have used 2-p-iodophenyl-3-p-nitrophenyl-5phenyltetrazolium chloride (INT) rather than TTC as the electron acceptor (e.g., Nannipieri et al., 1990). Inasmuch as the data derived from the procedures described above are obtained on the same subsamples of soil, removed after different


41

periods of incubation from the same master jars, on which CO, evolution data are being continuously obtained, changes in the metabolic activity, either gross or specific, depending on the substrates added, can be correlated with changes in various specific indicators of the microbiota (e.g., species diversity, antibiograms, biochemical and nutritional characteristics, enzyme activity, fate of the GEMs), both over time and between uninoculated soil and the soil inoculated with a GEM or the homologous parental strain. The redundancy purposely built into these experiments (e.g., CO, evolution and dehydrogenase activity) provides an internal control on the validity and sensitivity of the individual indicators. Furthermore, the degree of correlation between the various indicators should identify those assays that are clearly redundant and can be eliminated in the further development of a standard battery of assays with which to evaluate the potential impacts of any GEMs introduced into soil or other natural habitats on microbial populations and microbe-mediated ecological processes. F. NITROGENTRANSFORMATIONS

Studies on the effects of GEMs on specific biochemical transformations of fixed nitrogen are performed with a perfusion system that percolates nitrogen-containing water through a soil column under continuous recirculation. The perfusion apparatus recommended is a modification of that used by Macura and Stotzky (1980), Kunc and Stotzky (1980), and Bewley and Stotzky (1983~)(Fig. 5). A recent modification of this apparatus has reduced the cost and space requirements (Fig. 6). Soil, uninoculated or inoculated with either a GEM or the homologous parent, is amended (either directly or via the perfusion solution) with an ammonium salt, simple nitrogen-containing organics (e.g., amino acids), or complex nitrogen-containing organics (e.g., proteins or plant tissues), and net nitrification and nitrogen mineralization rates are determined. The perfusates are analyzed, after appropriate times, for different nitrogen fractions (i.e., a-amino, ammonium, nitrite, and nitrate nitrogen) and pH, and the soil can be analyzed for the presence of the GEMs or the novel genes, species diversity, and other parameters. Although nitrogen transformations can also be studied in subsamples of soils from the master jars, extraction of the various nitrogen fractions from soil is tedious, time-consuming, incomplete, and imprecise. In contrast, the soil perfusion technique is highly sensitive, easy to perform, and, as it is a continuous system, yields excellent kinetics. This technique has been used extensively and successfully to evaluate the

42

G . STOTZKY ET AL.

h y e r of fiberglass

-mm mesh opening)

Unless indicatedotherwise, all glass tubing is 6 mm O.D. Pyrex and all rubber tubing is 1/4 in. I.D. latex

FIG. 5. Perfusion unit for studying transformations of fixed nitrogen in soil (Jones et al., 1991).

43

RELEASE OF GENETICALLY ENGINEERED MICROORGANISMS 21em00

4

c*

FIG. 6. Modified perfusion unit for studying transformations of fixed nitrogen in soil (J. D. Doyle and G. Stotzky, unpublished).

44

G. STOTZKY ET AL.

effects of sulfur dioxide, acid precipitation, heavy metals, clay minerals, and GEMs on nitrogen transformations in soil (e.g., Bewley and Stotzky, 1983c;Jones et al., 1991; Kunc and Stotzky, 1980; Macura and Stotzky, 1980; see Stotzky, 1986). If significant changes in nitrogen transformations are observed as the result of inoculating a GEM, the kinetics of nitrogen transformations in subsamples of soil from the master jars should be compared to the kinetics observed in the perfusion technique, to verify the effects of the GEM. Transformations of fixed nitrogen will be of particular interest when GEMs containing nif genes are inoculated into soils, with and without energy and carbon sources (e.g., plant residues). If dinitrogen fixation is enhanced, especially in the presence of such sources, this could result in increased nitrification of the resultant ammonium, which could affect numerous microbe-mediated ecological processes as the result of the attendant accumulation of protons (Lee,decreases in the pH of the soil). The magnitude of these effects will depend, to a large extent, on the types and amounts of clay minerals present in the soils, as the buffering capacity of soils, which controls the magnitude of changes in pH, is strongly influenced by the clay mineralogy (e.g., Stotzky, 1972,1986). Procedure. Sieve moist soil through a 5-mm screen, collect on a 2-mm screen, and place 40 to 50 g of soil, oven-dry equivalent, in glass columns (180 x 25 mm, OD). Place a loose, thin (1-2 mm) layer of fiberglass (glass wool) at either end of the soil column. Place the bottom layer on a circular piece of nylon or plastic mesh (1mm) that is supported by a ring of polyvinyl chloride with four sections cut out of the bottom of the ring, which assists in maintaining the integrity and water flow characteristics of the column. Perfuse the soil with 200 ml of an amino acid solution or some other form of organic nitrogen (usually -140 pglml a-NH,+ N) from a reservoir (e.g., a 250-ml separatory funnel or a 200-ml specimen jar) that is connected to the perfusion column via glass and latex rubber tubing. Connect individual columns in series, via a manifold (see section on Metabolic Activity), to the vacuum source. Should a soil column collapse (an infrequent occurrence), insert short, narrow sections of Pasteur pipettes vertically through the soil to provide drainage, aeration, and proper vacuum conditions for continuous perfusion. Accomplish continuous perfusion by suction provided by a vacuum source (e.g., a pump or water aspirator). Equalize the perfusion rates of the individual columns by inserting a 25-gauge hypodermic needle into the outlet tube to the vacuum manifold. Adjust the flow rates of individual perfusion columns to 2 to 4 ml/min with screw clamps on the latex tubing of the air inlet. Measure the flow rate by placing a small


45

test tube (e.g., a Durham or microcentrifuge tube) below the perfusate inlet tube on the top of the glass column, and determine the amount of perfusate collected per unit time. Do not collect for more than 1 min, as perfusion will be disrupted, but repeat several times. Calibrate the number of drops per unit time falling from the perfusate inlet tube with the volume collected. Use the number of drops per unit time to verify flow rates during an experiment. Add GEMS or homologous parental strains at desired concentrations (e.g., lo7 CFU/g soil) to the top of the soil columns. Remove 5 to 7 ml of perfusate, with a pipette or syringe from each separatory funnel via the air inlet tube or by syringe directly from the specimen jar reservoir through the sampling port, periodically during the continuous perfusion of the columns (e.g., day -1, one day before the addition of the organic nitrogen source and inoculation; day 0 , 2-3 hours after addition and inoculation; and at different times after inoculation), and place in screw-capped test tubes. Replace the amount of perfusate removed with an equal amount (5-7 ml) of tap water. Within an hour after samples are collected, measure the pH, add 3 drops (-0.1 ml) of 0.1% HgCl,, and store the samples at 4°C until analysis of the different forms of nitrogen. Determine the concentrations of the different forms of nitrogen in the perfusates with a Technicon Autoanalyzer I1 (or other type of analyzer, or manually). Measure the concentration of a-NH,' N by reacting the sample with trinitrobenzene sulfonic acid in a buffered alkaline medium, heating at 65°C to produce a chromophore, and measuring the absorbance at 420 nm (Technicon Industrial Method No. 493-77A). Determine the concentration of NH,' N with the Berthelot reaction, which involves the formation of a green-colored compound, believed to be closely related to indophenol, and measure the absorbance at 630 nm (Technicon Industrial Method No. 98-70W/A). Determine the concentration of NO,- N plus NO,- N by using a cadmium reductor column that reduces NO,- to NO,- N. The NO,- N reacts with sulfanilamide under acidic conditions to form a diazo compound, which couples with N-1-naphthylenediamine dihydrochloride to form a reddish dye. Measure the absorbance at 520 nm (Technicon Industrial Method No. 100-70W/B).Determine the concentration of NO,- N separately, as for NO,- N plus NO,- N, except eliminate the reductor column (Technicon Industrial Method No. 103-70W/C).Subtract the concentration of NO,- N from the total NO,- N plus NO,- N content to determine the concentration of NO,- N. Collect the data as numerical peak heights (e.g., on a modular-digital printer), and substantiate by flatbed recorder graphs for each chemical analysis. Correct the raw data from the modular-digital printer of the Techni-

46

G. STOTZKY ET AL.

con Autoanalyzer 11 with the perfusate dilution factors and for the dilutions that were necessary for the samples to fall within the range of the standards for each of the chemical analyses. Correct for the reduction in the quantity of total nitrogen in the perfusate, as the result of the sequential sampling and replacement of perfusate with an equal amount of water, with appropriate dilution factors. For example, after removal of 7 ml on day 0,200 m1/193 ml = 1.036; after removal of 7 ml on day 1, 200 m1/186 ml = 1.075; after removal of 7 ml on day 2, 200 m1/179 ml = 1.117. The correction factor can also be calculated on the basis of the amount of added nitrogen removed at each sampling. For example, if 140 pg/rnl of a-NH,' N is added initially, a 200-ml reservoir will contain 28,000 pg of added nitrogen. After the removal of three sequential 7-ml samples, the amount of nitrogen remaining in the reservoir will be 25,161.7 pg [28,000 pg - 7(140 pg/ml) = 27,020 pg/200 ml or 135.1 pg/ml; 27,020 pg - 7(135.1 pg/ml) = 26,074.3 pg/200 ml or 130.37 pg/ml; 26,074.3 pg - 7(130.37 pgl ml) = 25,161.7 pg/2OO ml or 125.81pglml], and the dilution factor will be 140 pg/ml + 125.81 pglml = 1.113. There is a slight difference between dilution factors calculated by the two methods. However, if the same method is used consistently, the small difference between methods is irrelevant, as the critical comparisons in nitrification rates are between uninoculated soil and the soil inoculated with a GEM or the homologous parent. Adjust the NH,' N data to account for the contamination by a-NH,' N. In addition to analyzing NH,' N standards, analyze a series of corresponding a-NH,' N standards using the analytical procedure for NH,' N. Generate two separate standard curves and their respective equations. Subtract the corresponding a-NH,' N value from the NH,' N concentration determined in the assay, to obtain the corrected NH,' N Concentration. After determining the NH,' N concentration (including any contamination from a-NH2+N) from the standard curves, rearrange the equations to solve for absorbance as a function of N concentration, set the two equations equal to one another, and solve for NH,' N concentration as a function of the a-NH,+ N concentration. This provides a measure a-NH,' N that appears as NH,' N in the alkaline phenol assay.

G. NONSYMBIOTIC DINITROGEN FIXATION The potential effects of GEMS on N, fixation can be estimated by the acetylene (C,H,)-reduction technique (Hardy et al., 1968; Knowles, 1982). The technique quantifies the activity of the nitrogenase complex that is responsible for N, fixation. A factor of 3 moles of ethylene (CZH4)


47

formed per mole of nitrogen reduced is used to convert the CzH4 data to the amount of N, fixed. Procedure. Place 30 g of soil, oven-dry equivalent, at its -33-kPa water tension in 130-ml French square bottles or in the vials used in the metabolic studies, which serve as incubation chambers for the C,H,-reduction assay. Prepare the GEMS and the homologous parental strains in appropriate liquid media amended with appropriate selective factors to maintain culture integrity. Suggested controls include (I)tap water only; (2) low concentration (e.g., 7.7 x l o + atm) of CzH4 and tap water; and (3) a GEM or its homologous parental strain without C,H,. Mix the soils with the inoculum or an equivalent volume of tap water, add to the bottles or vials, and cover with rubber septa for the C,H, reduction assay. Between assays for N, fixation, place the uncovered bottles or vials in the master jars used for the CO, evolution studies, and continuously flush with water-saturated air. Perform C,H,-reduction assays at various times before and after the addition of the inoculum (e.g., day -1, one day before inoculation; day 0 , immediately after inoculation: and at different times after inoculation). Add C,H, (industrial grade) with a gas-tight syringe to the septa-fitted bottles after evacuation, with a syringe, of an equal volume of headspace air (e.g., 6.5 ml from a 130-ml bottle) to yield 0.05 atm. Incubate the bottles for 6 hours at 24 2 2°C in a water bath, remove two 20-pl gas samples with a gas-tight syringe from each bottle, and inject each sample into a gas chromatograph (e.g., Perkin-Elmer 3920) equipped with a flame-ionization detector and a 6-foot, 0.085-mm ID column packed with Porapak N (80-100 mesh). Maintain the temperatures of the injection chamber, oven, and detector at 100,60,and lOO"C, respectively, and the flow rates of the nitrogen carrier gas, compressed air, and hydrogen at 30, 300, and 30 ml/min at 60, 60, and 40 psi, respectively. Determine the concentrations of CzH4 formed by the reduction of C,H, from standard curves obtained with pure CzH4. Determine the background production of CzH4 (e.g., from the indigenous microbiota that was not related to reduction of C,H,) and any CzH4 released from deterioration of the septa using a control soil that received no C,H, or inocula. Correct the concentrations of C2H4 for background CzH4 production, according to the method of Knowles (1982). Use a control that received a small quantity of C,H4 (e.g., 7.7 x l o 4 atm; i.e., 10 plhottle) to detect the metabolism of endogenous C,H, by the indigenous soil microbiota. Acetylene inhibits the metabolism of CzH4, and the net accumulation of endogenous C2H4 could be greater in the presence of high concentrations of C,H, than in its absence. This CzH4 would be

48

G. STOTZKY ET AL.

measured as part of the CzH4 produced as a result of C,H, reduction. Background C,H, levels can be a problem when studying C,H, reduction by free-living nitrogen-fixing microbes in soil wherein the amount of C,H, reduction is usually low. Record the C,H, concentrations, corrected for background errors and soil water content, as nanomoles per gram soil, oven-dry equivalent, per hour. AND COMPETITIVE ABILITYOF H. GROWTHUTES GENETICALLY ENGINEERED MICROORGANISMS

The growth rates of GEMs and their homologous parental strains, their ability to compete with indigenous soil microbes, and the transfer of the novel genetic information can be evaluated by the soil replicaplating technique (Krasovsky and Stotzky, 1987; Rosenzweig and Stotzky, 1979, 1980; Stotzky, 1965b, 1972, 1973, 1986; Weinberg and Stotzky, 1972). These studies will provide data on the influence of introduced novel genes on the growth rates of the host bacteria in soil and on their ability to compete and react to amensalism. Procedure. Inoculate the GEMs or the homologous parental strains into the center of petri plates containing sterile soil adjusted to the - 33-kPa water tension, and inoculate representatives (isolated in the microbial assays) of the indigenous soil microbiota (e.g., bacteria, including actinomycetes, and fungi) into equidistant sites around the GEMs or homologous parents. Prepare the sterile soil plates by placing 40 to 50 g of soil, oven-dry equivalent, at the - 33-kPa water potential plus about 2 ml of water (which will be lost during autoclaving; the amount of additional water to add will depend on the soil used) into thick-walled glass petri plates. Autoclave the plates for 20 min at 121OC,cool to mom temperature, and autoclave again for 20 min. Check for sterility, as described earlier. Place the covered sterile soil plates in a humidified chamber for 1week, to allow for dissipation of any toxic compounds. After inoculation, incubate the soil plates in a high humidity chamber under fluctuating temperatures (e.g., 23 to 27OC), to prevent desiccation. Replicate from the soil plates to selective agars periodically (e.g., every 4 days) with a replicator constructed from acrylic plastic and stainless steel nails (20 gauge, 3.8 cm long) (see Stotzky, 1965b, 1973, for details on construction and use) that are sterilized with alcohol and flaming. The design of the replicator permits numerous replications from the same soil plate without significant disturbance of the soil. Change the design of the replicator (i.e., placement of the nails) to accommodate studies of physiological groups, antibiotic-resistant phenotypes, and biochemical characteristics, as described above. Re-


49

cord the growth of all inoculated organisms on maps of the soil plates. Calculate growth rates (in mmlday). If consistent and significant differences in growth and competitive ability between the GEMs and the homologous parents are observed, inoculate the GEMs or the homologous parents into the center of plates containing nonsterile soil, and replicate to cycloheximide-containing media highly selective for the GEMs, the novel genes, and the homologous parental strains (e.g., agars containing nalidixic acid), to prevent overgrowth by the indigenous soil microbiota. If necessary, confirm the presence of the novel genes by DNA probes and other molecular techniques, as described above (e.g., Stotzky et a]., 1990).If the occurrence of amensalism is suggested from these studies in soil, streak the presumed producers of the amensalistic substance(s) against the GEMs and the homologous parents on agar (e.g., Rosenzweig and Stotzky, 1979,1980).

I. STATISTICAL DESIGNAND ANALYSIS Considerable redundancy should be designed into the studies [e.g., various measures of heterotrophic microbial activity in the master jar studies; rates of heterotrophic deamination in the perfusion studies and heterotrophic metabolic activity in the CO, evolution studies; rates of nitrification in the perfusion studies and numbers of nitrifiers (and denitrifiers) in the master jar studies; survival of the GEMs in the master jar and perfusion studies and their growth and competition in the soil replica-plating studies], This redundancy not only provides internal controls for the validity and sensitivity of the different assays, but it should indicate those assays that best reflect the effect of an introduced GEM on microbe-mediated ecological processes in soil. All data should be statistically analyzed, using the appropriate types of analyses (e.g., Steel and Torrie, 1980).The design of the studies must be predicated on the types of statistical analyses that will be applied to the data. For most studies, the data can be expressed as the mean & the standard error of the mean, and the means should be compared by the two-tailed Student’s t-test. Wherever possible or necessary, analysis of variance and regression analyses, including the Duncan Multiple Range or the Tukey Honestly Significant Difference tests, should be used. These simple analyses have been successfully applied in previous studies in our laboratory on the effects of chemical and physical environmental perturbations on microbes and their activities in soil (e.g., Stotzky, 1986).Nevertheless, it is advisable to consult a statistician before the design of experiments. Moreover, the relation between statistical significance and ecological

50

G.STOTZKY ET AL.

significance must be considered when evaluating the data. For example, differences in some microbial populations and processes in a soil inoculated with a GEM or with the homologous parent without the novel gene may be statistically significant in some samplings of an experiment but not on other sampling days, and the ecological significance of such differences is, at this stage of development, a matter of judgment on the part of the investigator. This is, obviously, an area that requires extensive study. Nevertheless, even at this stage of development, a GEM or its homologous parent can be assumed to have a significant ecological effect when more than one parameter of microbial populations and microbe-mediated processes is affected and the effects on these parameters occur consistently over time. The concepts and techniques of the ecologic dose (EcD) quantification developed for assessing the impacts of environmental toxicants on microbe-mediated ecological processes might be applicable to express the effects of GEMs on these processes in soil and other natural habitats (Babich and Stotzky, 1983,1985;Babich et a].,1983).The EcD quantification is defined as the dose of a toxicant that decreases a specific microbe-mediated ecological process by a certain percentage. This quantification enables the identification of “high risk” and “low risk” environments, namely, environments in which the effects of a toxicant are magnified or reduced, respectively. The quantification is also helpful in the evaluation of the physicochemical characteristics of an environment that are most important in mediating the activity of a toxicant, which might suggest methods with which to ameliorate its toxicity in habitats that are already contaminated and, thereby, enable their reclamation. Although an inanimate toxicant is different from a living GEM, especially in terms of containment, there are numerous similarities when these foreign entities are introduced into natural habitats. Consequently, the application of the concepts and techniques of the EcD quantification, with appropriate modifications, might provide insights into the potential impacts of GEMs on microbe-mediated ecological processes in soil and other natural habitats. III. Representative Results

To illustrate the applicability of these techniques to the study of the potential impacts of the introduction of GEMs on microbial populations and microbe-mediated ecological processes in soil, some representative results are presented. These studies were conducted both in the Laboratory of Microbial Ecology of the Department of Biology at New York University and at the Corvallis Environmental Research Laboratory of the U.S. Environmental Protection Agency, to provide


51

TABLE IV PHYSICOCHEMICAL CHARACTERISTICS OF THE ITCHAWAN AND MILLICAN SOILS Measure

Kitchawan soil.

Millican soilb

Sand, % Silt, % Clay, % Organic matter (loss on ignition), % Total nitrogen, Yo PHw Cation-exchange capacity, cmollkg ExchangeableH, cmol/kg Bulk density (disturbed core), mg/m3 Soil water content (at - 33 H a ) , % Phosphorus (available),mg/kg Potassium (available),mg/kg Magnesium (available),mglkg Calcium (available],mg/kg Iron (available),mg/kg Aluminum (available),mglkg Manganese (available),mg/kg Zinc [available], mg/kg Copper (available),mglkg

56.8 33.8 9.4 2.62 0.13 4.9 11.9 11.9 1.13 17.4 2.6 26.0 17.6 105.0 7.9 180.9 10.1 3.5 0.4

68.1 29.4 2.5 1.95 0.06 5.9 13.2 4.0 1.27 11.1 8.6 286.0 215.8 981.0 1.1 16.0 12.1 0.4 0.4

0 Provided by the Agronomy Analytical Laboratory of Cornell University. Selected analyses were confirmed by the College of Environmental Science and Forestry, State University of New York, Syracuse. b Provided by the Agronomy Analytical Laboratory of Cornell University,Oregon State University Soil Testing Laboratory, and the College of Environmental Science and Forestry, State University of New York,Syracuse. Analyses from the different laboratories varied slightly for individual parameters, and the means of the analysis are indicated.

interlaboratory validation of the techniques. In New York, a sandy loam soil collected at the edge of a plot containing young magnolia trees at the Kitchawan Research Laboratory of the Brooklyn Botanic Garden in Ossining, New York, was used. This soil has been extensively used in previous studies on the effects of chemical and physical perturbations on microbe-mediated ecological processes, as well as on the survival of and gene transfer by GEMS. Consequently, there is a large database available on the Kitchawan soil (e.g., Devanas et a]., 1986; Stotzky, 1986;Stotzky et al., 1990). In some studies, the Kitchawan soil was amended with different concentrations of the clay minerals montmorillonite or kaolinite. In Corvallis, a xeric sandy loam soil from the Millican Limited Use Area in central Oregon was used. Some physicochemical properties of the Kitchawan and Millican soils are presented in Table IV. The GEMS and the homologous, plasmidless parental strains used, as

52

G. STOTZKY ET AL. TABLE V

GENETICALLYENGINEERED MICROORGANISMS AND HOMOLOGOUS PARENTAL STRAINS AND THEIR MAINTENANCE AND ISOLATION MEDIA"

GEM Species

Strain

Parent Phenotype

108(R388::Tn1721)b Nx: Tp', Tcr W3110(R702)d Tm+,Km', Smr, Su', Tcr,Hgr Tra+, Apt, Nm*, J53(RP4Id Escherichia coli Tcr,Kmr Pseudomonas putida PP0301(pR0103)f Nxr,Tc'. Hg', degrades 2,4-D to chloromaleylacetate

Enterobacter cloacae Escherichia coli

Strain

Phenotype

107c W 3 110

Nxr Prototrophic

J53'

Pro-, Met-

PP0301s Nxr

L agar, Luria agar; Nx, nalidixic acid: Tc, tetracycline: Tp. trimethoprim; MAC, MacConkey agar: TNA, tryptone, ysast extract, dextrose, and NaCl agar. Subscripts indicate concentrations of the antibiotics (clglml of medium]. See Table In for abbreviations for antibiotics to delineate phenotypes. L war + Nxsoo + Tc,~+ Tpm. Lagar + Nxm. MAC TcZ~. 0 MAC. f TNA + N X ~ W +Tc~. 8 TNA + N X ~ .

+

well as their maintenance and isolation media, are presented in Table V. Studies with each GEM and homologous parent were repeated numerous times in both the Kitchawan and Millican soils. Consequently, the data are extensive and repetitious, and only representative data for each soil are presented for the purpose of demonstrating the applicability of the techniques. Greater details on these studies and their implications can be found elsewhere (e.g.,Doyle et al., 1991;Jones et al., 1991;Short et al., 1991). A. METABOLIC ACTIVITY(CARBON DIOXIDEEVOLUTION)

The basal rate of metabolism of both soils, whether uninoculated or inoculated with a GEM or a homologous parent, was low. Consequently, the soils were amended with glucose (I%, w/w) at the beginning of the studies. In some experiments, glucose (1% w/w) was also added (pulse, P) during the incubation. For the Kitchawan soil, respiration data are presented for four GEMs and their homologous parents from two representative studies in which two GEMs and their respective parents were evaluated in both studies, to provide an indication of the reproducibility between studies.


53

The highest rate of CO, evolution occurred within the first 2 days of the incubation (Figs. 7 and 8 ) . In general, soils inoculated with a GEM or a homologous parent had slightly higher rates of respiration than the uninoculated control soils during this early period, probably as the result of the addition of the inocula (106 to lo7 CFU/g soil), which were in the log phase. In contrast, the uninoculated control soils usually had slightly higher rates of respiration later in the incubation (days 4 and 5), probably as the result of higher levels of residual glucose than in the inoculated soils. After the peak in CO, evolution and when most of the added glucose had been mineralized or incorporated into biomass, the rates of respiration decreased to a stable, basal level. There were no consistent and persistent effects on soil respiration that could be attributed to the introduction of the GEMS or their homologous plasmidless parents. The increases in the rate of respiration of the Kitchawan soil pulsed with glucose after 14 days of incubation were similar for all inoculum treatments and also showed no effects that could be related to the introduced GEMs or their homologous parents. The lower rates of respiration of the uninoculated control soils were no longer apparent, indicating that the slight effects of adding GEMs or their parents noted during the first week of incubation were transient. There were differences in rates of respiration between the first and second additions of glucose: the highest rate of respiration occurred within the first 2 days after the initial addition of glucose, whereas it occurred 3 to 4 days after the glucose pulse. Moreover, the amount of CO, evolved was greater after the pulse than after the initial addition of glucose, However, these differences were independent of the addition of the GEMSor their homologous parents. Furthermore, the general pattern of respiration was similar in all studies conducted with the Kitchawan soil.

B . SPECIES DIVERSITY The data for species diversity in the Kitchawan soil are presented as histograms, with error bars indicating the standard errors of the means and letters indicating the Duncan Multiple Range values, as three GEMS and their homologous, plasmidless parents were evaluated simultaneously. For the data from the Millican soil from Oregon, wherein only one GEM and its homologous parent were evaluated in each experiment, line graphs are used, with the standard errors of the means indicated when they were larger than the dimension of the symbols, and the Tukey values were tabulated separately. The best methods for pre-

Daily respiration

8o

1

60

Pa3 8

40

5

El 20

OL Cumulative respiratlon

500

P

-

400

8

300

El

200

5

control

Entemlmcterdoacae107 Entetvbactterdaecae lOB(R388) EscherlchiawllJ53 Esdmrkhia wli&3(RP4) E8Ch8rkhia wli W3110 Eschotkhia wll w31 lO(R702)

100

0

0

10

20

30

40

Day

FIG.7. Daily and cumulative rates of CO1 evolution from Kitchawan soil inoculated with Enterobacter cloacae 108(R388), Escherichia coli J53(RP4). Escherichia coli W3110(R702), or the homologous plasmidless parents E. cloacae 107, E. coli J53, or E. coli W3110, or not inoculated.The soil was amended on day 0 with 1%(w/w) glucose and pulsed with 1%glucose on day 14. 54

ioa

Daily respiralion

80

60

40

20

0 600

-

Curnulathre respiration

500.

-c

400.

control Entembauerdoacao 107

300-

200.

i

0

10

20

30

40

Day

FIG. 8. Daily and cumulative rates of CO, evolution from Kitchawan soil inoculated with Enterobacter cloacae lO8(R388), Pseudomonas putida PP0301(pRO103), Escherichia coli W3110(R702), or the homologous plasmidless parents E. cloacae 107, P. putido PP0301, or E. coli W3110, or not inoculated. The soil was amended an day 0 with 1% (w/w) glucose and pulsed with 1% glucose on day 14. 55

56

G.STOTZKY ET AL.

senting the data from these extensive and complex experiments have not been identified and await comments and suggestions from peer reviewers. The numbers of total bacteria in the Kitchawan soil remained essentially constant throughout the incubation (Fig. 9). The addition of glucose, either initially or after 14 days, slightly increased the numbers of total bacteria, especially after the glucose pulse. However, there were no consistently significant differences that could be attributed to the introduction of the GEMs or their homologous parents. Spore-forming bacteria comprised about 10% of the total bacterial population in this soil (Fig. 10).The initial addition of glucose resulted in a slight increase in spore-forming bacteria, whereas the glucose pulse resulted in a decrease in numbers. However, as with total bacteria, the changes with time in the numbers of spore-forming bacteria were not correlated with the inoculation of the soil with either the GEMs or their respective parents. The numbers of fungal propagules remained relatively constant, regardless of whether the soil was pulsed with glucose on day 14, and there were no consistent differences that could be attributed to the GEMs or their parents (Fig. 11). The numbers of cellulose-utilizing bacteria increased significantly during the second week of incubation (Fig. 121,presumably as the result of the depletion of the added glucose, which provided the microbiota with a readily available source of carbon and energy. However, the pulse of glucose on day 14 did not reduce the numbers of celluloseutilizing bacteria. Inasmuch as the numbers of total bacteria remained essentially constant throughout the incubation (Fig. 9),these data indicated that a change occurred in the diversity of the bacterial population with time. Concomitant with the general increase in the numbers of cellulose-utilizing bacteria, there was a trend to higher numbers in soil inoculated with the homologous parent than with the respective GEM or not inoculated, especially after 14 days, irrespective of the glucose pulse. The ecological significance of this trend is not known. Chitinoclastic bacteria, predominantly actinomycetes, exhibited the same general trends as the cellulolytic bacteria. The numbers of chitinutilizing bacteria increased significantly between days 3 and 8 and again after day 14 (Fig. 13). As with the cellulolytic bacteria, there was a general trend to slightly higher numbers of chitinoclastic bacteria in soil inoculated with the homologous parental strain than with the respective GEM or not inoculated. However, these differences were usually small and probably not ecologically significant,

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a

Control (NOaddition) E.coliW3110 E. coli w 3 i i q ~ 7 0 2 1

~.putida~~0301 P. putida PP0301[pRO103] €.cloacae107 E. doacae 1Oa[R3as]

-r

-.I

0

3

8

14

-r

28

-r

P+7

P+21

Days FIG. 9. Numbers of total heterotrophic bacteria enumerated in Kitchawan soil inoculated with Enterobacter cloacae 108(R388), Pseudomonas putida PPO301(pRO103),Escherichia coli W3110(R702), or the homologous plasmidless parents E. cloacae 107, P. putida PP0301, or E. coli W3110, or not inoculated. The soil was amended on day 0 with 1% (w/w) glucose and pulsed (P) with 1% glucose on day 14. The standard errors of the means and the Duncan Multiple Range values are indicated. Data are shown for soil amended with glucose only on day 0 (days 0 through 2 8 ) and for soil pulsed with glucose on day 14 (i.e., 21 and 35 days after the initial amendment with glucose on day 0 ) .

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control (Noadditkn) E. cdi W3110 E.coli W311O(R702j P.puhldapp0301 P. pub;da PPo301[pROlO

E.doacae 107

-r

T

3

8

14

28

.

-a

P+7

P+21

Days FIG. 10. Numbers of spore-forming bacteria enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

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control (NO addion) E.wliW3110

w

E. wli W3110[R'102] P.putidaPp0301

w

P. putida P P O ~ O I I ~ R O ~ O ~ J

E.doacae 107 ~.doacael~mj

-r

0

3

-r

8

.r

14

-r

28

P+7

P+21

Days FIG.11. Numbers of fungal propagules enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

8

w H w

e d e

Control (No addition) E.COI~W~IIO E. coli W3110[R702]

c d c

P.putidaPp0301 P. putida PPO301[PRO103] E.doacael07 E.cloacaeIWR3881

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0

3

0

r

03 0

2

0

-r

T

8

-l

14

T

20

P+7

P+21

Days FIG. 12. Numbers of cellulose-utilizing bacteria enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

IControl (No addition) E. coli W3110

E. coli W3110[R702] d

P. putida PP0301 P. putida PP0301IpRO103j

E. doacae 107

a

e .

d

I

cbc ab

a

dcdbc h a

e

C

a

e

aaaaaaa

a

E. cloacae 108[R388]

aabaaaa

-r

1

0

3

7

8

-r

1

14

28

P+7

P+21

Days FIG. 13. Numbers of chitin-utilizing bacteria enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

62

G. STOTZKY ET AL.

The numbers of denitrifying (Fig. 14) and nitrate-reducing (Fig. 15) bacteria remained essentially constant in both studies during the initial 14 days of incubation and then increased substantially after 14 days in both the glucose-pulsed and nonpulsed soil. However, there were no consistent trends attributable to inoculation of the soil with either the GEMS or their respective parents. The numbers of ammonium oxidizers decreased during the first 2 weeks of the incubation and then increased by about one order of magnitude, regardless of whether the soil was pulsed with glucose, to the levels originally present (Fig. 16). These fluctuations in numbers were independent of the addition of the GEMs or their respective parents. Some fluctuations over time were also observed in the numbers of nitrite oxidizers (Fig. 17). However, these differences could not be related to the presence of the GEMs or their parents. The numbers of protozoa increased by one to two orders of magnitude during the incubation (Fig. 18). Similar to the trends observed with the cellulolytic and chitinoclastic bacteria, there was a trend to higher numbers of protozoa in soil inoculated with the homologous parental strains than with the respective GEMS or in the uninoculated soil. However, this trend was not consistent, and the variability among replicates of the same treatment was often higher than with other groups of organisms, suggesting that this trend was not ecologically significant.

c. ACTIVITYOF S O I L ENZYMES Representative data of the activities of selected soil enzymes are also presented as histograms from studies in glucose-pulsed Kitchawan soil. The dehydrogenase activity fluctuated during the incubation (Fig. 19). The activity increased after the initial addition of glucose but decreased after the glucose pulse on day 14. ,Despite the substantial variability in dehydrogenase activity with time, there were no consistent differences that were attributable to the introduction of the GEMS or their respective parents. There were no apparent relations between dehydrogenase activity and respiration rates (Fig. 8), even though dehydrogenase activity is considered by some investigators (e.g.,Nannipieri et al., 1990) to be a measure of metabolic activity. Although there were increases in both dehydrogenase activity and CO, evolution after the initial addition of glucose, which probably reflected a greater metabolic activity as a result of the introduction of both the bacteria and glucose, the addition of glucose on day 14 resulted in a decrease in dehydrogenase activity, whereas it resulted in an increase in CO, evolution.

Control (NO addition)

a

E.cdiW3110

v

E. coli W3110[R702] P.pufidaPPO301

aaaaaaa

aaaaaaa

,

b

C

P. putida PPo3o1[pRo103]

a

ab

a

T.

E.doacael07

T

aaaaaaa

T

.r

T

0

3

.

8

.r

14

28

Pi7

P+21

Days FIG. 14. Numbers of denitrifying bacteria enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated (Joneset al., 1991). See Fig. 9 for details.

w

Control (No addition)

0

E.CQI~W~IIO E. coli W311qR702j

€l P . p u t i d a P W 1 aaaaaaa

P. putida PPO3011pRO1a3j E.doacaelO7

rT

T

-

E.doacae100FU001

I aaaaaa

a

T

aaaaaaa

aaaaaaa

T

-r

0

-r

7

3

0

14

-

20

P+7

.

P+21

Days FIG.15. Numbers of nitrate-reducing bacteria enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated (Joneset al., 1991).See Fig. 9 for details.

aaaaaaa

aaaaaaa T

aaaaaaa T I

aaaaaaa

T

aaaaaaa Control (NOaddition) E.coliW3110 E. cdi W31 lO[R702]

E

P. puma PP0301 P. putida PP0301[pR0103] E. cloacae 107 E. cloacae 108[R368]

-r

0

3

Y

-I-

8

14

28

P+7

P+21

Days FIG. 16. Numbers of ammonium-oxidizing bacteria enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated (Joneset al., 1991). See Fig. 9 for details.

--

I

aaaaaaa a

aaaaaab

T

aaaaaaa

I

aaaaaaa

aaaaaaa

3

8

T

b*

m m

-..

-r

0

14

T

28

P+7

P+21

Days FIG. 17. Numbers of nitrite-oxidizing bacteria enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated (Joneset d.,1991). See Fig. 9 for details.


a

E.coliW3110

a

E. coli W3110[R702]

H

a

~.putida~~0301 P. puWa PP0301IpRO103]

E.doacae 107 E. cloacae 108[R388]

-r

-r

0

3

8

14

28

P+7

P+21

Days FIG.18. Numbers of protozoa enumerated in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

ab

3000 1

ab

.0

m

*

2000

P

a

abT

E

a) z 0

abc abc a ah

. cn


LL

El

n I-

m

1000

E . coli W31i0

-

E.coli W3110[R702]

=L

P. putjda PP0301 P. putida P P O ~ O I [ ~ R OI 031

E. cloacae 107 E. cloacae 10B[R388]

T

0

7

3

14

28

Pt21

Days

FIG.19. Dzhydrojienass ar.tivitv i n Kitc:hawan soil inoculated parents or not inoculatcd. Scc Fig. 9 for delails.

with three GEMS or their hnmologous plasrnidiexx


69

The acid phosphatase activity also fluctuated and decreased after both the initial and subsequent additions of glucose and then increased [Fig. 20). The initial addition of glucose, as well as the glucose pulse, also resulted in a decrease in the activity of alkaline phosphatase (Fig. 21). The activities of alkaline phosphatase were significantly lower than those of acid phosphatase (Fig. 20), which was probably a reflection of the low pH of the Kitchawan soil. However, despite the differences in the activities of the acid and alkaline phosphatases, there I were no consistent differences in the activities of either phosphatase that could be correlated with the addition of the GEMs or their respective parents. The activity of arylsulfatase was the lowest of the four enzymes evaluated [Fig. 22). The addition of glucose, both initially and on day 14, depressed sulfatase activity. There was a marked increased in activity by day 14, but there were no consistent trends in sulfatase activity that could be attributed to the addition of the GEMs or their homologous parental strains.

D. PH Although there were fluctuations in the pH of the Kitchawan soil during the 35-day incubation, these fluctuations were not related to the introduction of the GEMS or their respective homologous parental strains (Fig. 23). E. EFFECTOF ADDING 2,4-DICHLOROPHENOXYACETATEAND A GENETICALLY ENGINEERED MICROORGANISM CAPABLEOF ITS CATABOLISM ON MICROBIAL POPULATIONS AND PROCESSES IN SOIL

The data briefly described above were obtained in studies in which the GEMs or their homologous plasmidless parents, which served as internal controls for the novel genetic information, were added to soil without the concomitant addition of the antibiotics or heavy metals to which the novel genes conferred resistance or of the substrates for the enzymes encoded by the novel catabolic genes. Consequently, the absence of the stressors or substrates provided no ecological advantage to the GEMs. Moreover, there were no consistent or persistent statistically or ecologically significant effects on microbial populations and their processes as the result of the addition of the GEMs. To determine whether the addition of a substrate that a GEM, but not its homologous parent, could catabolize would result in an effect on microbial populations or their processes, a GEM [P. putida

Control(Noaddltion) E.coliW3110

E. CaH W31lqR7021 P.putidapp0301

P. put& PP0301IpR01031

a

a

-E.doacae 107

a a a aa'a a T

aaaaaaa

E.doacae

aaaaa aa

v

0

0

3

8

14

28

P+7

P+21

-

-Days FIG.$0. Acid phosphatase activity in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

aaaaaaa

P P

aaaaaaa

_Bab

T

aaaaaaa

Ti

~

aaaaaaa aaaaaaa

-

(No addition)

E.cdiW3110

E. wli W31 lO[R702]

E

~.putida~~0301

P. putida PP0301IpR0103]

E.doacae 107

1

E. doacae 1qR388] 3

-r

-r

7

0

8

14

Days

-r

28

-r

P+7

P+21

FIG.2'1. Alkaline phosphatase activity in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

aaaaaaa

aaaaaaa

aaaaaaa

T

7 [ ab

aaaaaaa aaaaaaa T U

N

.r

0

-r

3

-r

-r

8

14

-r

28

P+7

P+21

Days FIG.22. Arylsulfatase activity in Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

aaaaaaa

6

t

abab

a

b

ababab

aaaaaaa 5

bbbabbb 4

I

cL3

Control (No addiion) E.coliW3110

E. coli W3110[R702] 2

E ~.putida~~0301 P. putida PP0301IpR0103] E. doacae 107

E. doacae 108[R388] 1

0

7

0

3

-f

0

14

20

P+7

P+21

Days FIG.23. The pH of Kitchawan soil inoculated with three GEMS or their homologous plasmidless parents or not inoculated. See Fig. 9 for details.

74

G. STOTZKY ET AL.

PP0301(pR0103)]or its homologous parental strain (P. putida PP0301) 2,4was added to soil with the substrate (2,4-dichlorophenoxyacetate; D) on which the enzymes encoded by the introduced novel genes function. Pseudoyonas putida PP0301(pRO103) constitutively degrades 2,4-D to 2-chloromaleylacetate,(2-CMA), but it does not degrade 2,4-D to CO,, as the plasmid does not express chloromaleylacetate reductase. This construct was purposely chosen, as it represents, in many ways, an ideal GEM: it is genetically engineered to degrade a toxic xenobiotic, 2,4-D, to a harmless intermediate, 2-CMA, which is readily mineralized by the indigenous soil microbiota; and as the GEM derives essentially no energy from the transformation of 2,4-D to 2-CMA, it has no ecological advantage in soil, and once the 2,4-D has been degraded, the GEM will probably eventually disappear. The simultaneous addition of this GEM and 2,4-D to soil resulted in spme unanticipated effects. These effects are briefly described; more detailed information can be found in Doyle et al. (1991) and Short et al. (1991).The xeric Millican soil from Oregon was used in these studies, as the microbiota in this soil does not detectably metabolize 2,4-D. The soil was either not amended or amended with glucose (1%w/w), 500 pg 2,4-D per gram soil (500 ppm), or 1% glucose plus 500 ppm 2,4D and inoculated with P. putida PP0301(pR0103) or P. putida PPo301 to yield lo6 to 10' CFU/g soil, oven-dry equivalent, or not inoculated. Soil inoculated with the GEM and amended with glucose plus 2,4-D evolved significantly less CO, during the first 35 days of incubation than did inoculated soil amended with only glucose (Fig. 24). This reduction in the rate of respiration did not occur in $oil amended with glucose plus 2,4-D and either inoculated with the homologous plasmidless parent or not inoculated. There was no significant effect of the GEM on the rates of CO, evolution in the absence of 2,4-D, as the rates were the same as in uninoculated soil or in soil inoculated with the parent. After 35 days, the total amount of CO, evolved was essentially the same from all soil samples amended with glucose, with and without 2,4-D, and regardless of the inoculum, indicating that the inhibitory effects of 2,4-D in the presence of the GEM were relieved by this time. The numbers of fungal propagules in soil amended with 2,4-D and inoculated with the GEM decreased to undetectable levels after 10 days of incubation (Fig. 25). In unamended soil inoculated with the GEM, fungal propagules were not detectable after 39 days. Comparable decreases were not observed in soil inoculated with the parental strain or not inoculated and either amended or not with 2,4-D. The activity of dehydrogenases was stimulated by the addition of glucose, but it was inhibited by the addition of 2,4-D, either with or without glucose (Fig. 26). However, in soil amended with glucose plus

Glucose 180

I35 ,

.

Uninoculated

90

45

0 I80 c c

0

Ln

135

CT,

0 0'

-

PP030 1

90

\

c

0

P L

m

45

u

z o L

I80

135

Y'

.2,4-D1

90

103) 2,4-D

45

Unamended 0 0

5

10

I 5

20

25

30

3.5

40

45

50

Day FIG. 24. Cumulative rate of COz evolution from Millican soil inoculated with Pseudomonas putida PP0301(pRO103) or the homologous plasmidless parent P. putida PP0301 or not inoculated. The soil was either unamended or amended on day 0 with 1%(w/w) glucose, 500 ppm 2,4-dichlorophenoxyacetate(2,4-D),or glucose plus 2,4-D (Doyle et al., 1991). The standard errors of the means are indicated when larger than the dimensions of the symbols.

G.STOTZKY ET AL.

76

Uninoculatrd

' 1

I

7

6

S

8-

PP030 1 (PRO 103)

GI ucose

7-

l 6-

4

0

10

I

I

IS

20

25

30

SS

I

I

40

45

--I

SO

FIG. 25. Numbers of fungal propagules enumerated in Millican soil (Doyle et al., 1991). See Fig. 24 for details.

77

RELEASE OF GENETICALLY ENGINEERED MICROORGANISMS 1,5

1

Unlnocula t e d Glucose

1 .o

0.5 c

0.0 1 .f

1 .o

b c

a C

0.5

nn "."

1.5

. PP030 1 (PRO 103)

I .o

0.5

0.0

0

5

I0

15

20

25

50

55

40

41

SO

Day FIG. 26. Dehydrogenase activity in Millican soil (Doyle et al., 1991). See Fig. 24 for details.

78

G.STQTZKY ET AL.

2,4-D and inoculated with the GEM, but not in uninoculated soil or in soil inoculated with the parent, the suppression of dehydrogenase activity was relieved after 20 days as 2,4-D was degraded by the GEM. There were no consistent or persistent effects of the GEM o r of 2,4-D, either added alone or together, on the numbers of total heterotrophic, spore-formiqg, and chitinoclastic bacteria or on the activitied of alkaline and acid phosphatases or of sulfatases (data not shown; see Doyle ; et a ~ 'i991). In soil amended with 2,4-D and inoculated with the GEM, the concentration of 2,d-D, as determined by high-performance liquid chromatography+and gas chromatography-mass spectroscopy, decreased rapidly during the first 10 days of incubation to less than 200 ppm and then decreased more slowly to less than 100 ppm by day 53 (Short et al., 19911. Concomitant with the degradation of 2,4-D, the concentration of 2,4-dichlorophenol (2,4-DCP),the first degradation product of 2,4-D, accumulated t a greater than 70 ppm until day 38, after which time the concentration decreased. The accumulation of 2,4-DCP was apparently responsible for the reduction in the rate of CO, evolution and in b number of fungal propagules in soil amended with 2,4-D and inoculated with the GEM. In contrast, it was apparently the conversion of 2,4-D to 2,4-DCP that relieved the inhibition of dehydrogenase activity: Studies in pure culture with five fungal isolates from the Millican soiI"shciwedthat the growth of ond isolate'was completeljr inhibited by 10 ,ppm 2,4-DCP and that 5Q,ppm 2,4-DCP completely inhibited the k o h h of the other four. isolates, whereas even 200 ppm 2,4-D only reduced the growth of, the isolates (Short et a]., 1991). In sterile soil, 50 ppm 2,4;DCP reduced the spread (as measured by the soil replicaplating technique) of the fungal isolates by 90 to 99%, and ioo ppm completely inhibited the spread.

TRANSP~RMATIONS F. NITROGEN Studies on the .transformation of fixed nitrogen were Conducted with the Kitchawan soil, using the perfusion apparatus shown in Fig. 5. The soil was-inoculated with tbe GEMS and the respective homologous, plasmidless pareha1 strains (-4.5 x 10' CFU/g soil, oven-dry equivalent] and perfused with 200 ml of water to which glycine (140 pg aNH,' N/ml) had been added h day 0. After various periods of continuous perfusion, 7 ml of perfusate was removed and replaced with 7 ml of water, and'the perfusate samples were analyzed for a-NH,+, NH4+, NOz-, and NO,- N and pH.


-NHZ-

-

Control

(No addition)

NH4+

N03-

*..................

0

79

10

20

..........* ............*

30

40

50

Days

FIG. 27. Kinetics of nitrogen transformation in Kitchawan soil amended with 6% montmorillonite and not inoculated (Jones et al., 1991). The data in Figs. 27 through 31 are from experiments conducted concurrently.

The kinetics of nitrogen transformations in the Kitchawan soil amended with 6% montmorillonite and inoculated with each of the four GEMS or their homologous plasmidless parents were generally comparable to those in the uninoculated soil (Figs: 27-31). The changes in the various nitrogen fractions with time were corroborated by the changes in pH: during the ammonification stage, th'e pH increased, whereas the pH decreased as NO,- accumulated. The only exception occurred in soils inoculated with the strains df E. cloacae: the rate of nitrification and the amount of NO,- produced were greater in the presence of the parental strain than of the respective GEM (Fig. 31). However, these differences were not considered to' be large enough to constitute an ecologically significant effect (Jones et al., 1991). When the E. cloacae strains were added to the Kitchawan soil amended with different amounts of montmorillonite, the rate of formation of NO,- was greater in uninoculated soil than in soil inoculated with either the plasmid-containing or plasmidless strain in unamended soil (0% montmorillonite) (Fig. 32) and in soil amended with 9% montmorillonite (Fig. 33) but not in soil amended with 12% montmorillonite, wherein the rates were similar (Fig. 34). The most pronounced effect on nitrification resulted from the amount of montmorillonite added: as the clay content was increased, the lag phase of nitrification decreased and the rate of nitrification in-

80

G. STOTZKY ET AL. 200

---*--NH2-

+/:-I

--C-

E. coh W3110[R702]

NH4+

.......0.....

.......* ‘OTi

Z

7.5

..... \F””’ ..........*..

*...................7w

5

.........*

2.5 0

10

0

-0-

-E

.

20

E.

NH2-

COh

Days

30

40

50

W31 10

NH4+

N03-

a iw

lo

Z

Ti

1.5 5

.................**..........r

“8

2.5 0 0

10

20

30

40

50

Days

FIG. 28. Kinetics of nitrogen transformation in Kitchawan soil amended with 6% montmorillonite and inoculated with Escherichia coli W3110(R702)or the homologous plasmidless parent E. coli W3110 (Joneset al., 1991).

RELEASE OF GENETICALLY ENGINEERED MICROORGANISMS mn ---

.

-

---*--NH2........ 0...

81

E. coli J53[RP4]

NH4+ N02.

-

E loo a

'0%

z

7.5 5

...................&", ............... * .................*

2.5 0 0

10

30

20

40

50

Days

I-

---*--NH2-

-""

-

E. coli J53

N W

N03-

lo% 7.5 5

-.*...................t""'..............*........""L

2.5

0

10

2o

Days

30

40

50

FIG. 29. Kinetics of nitrogen transformation in Kitchawan soil amended with 6% montmorillonite and inoculated with Escherichia coli J53(W4)or the homologous plasmidless parent E. coli J53 (Joneset al., 1991).

82

G . STOTZKY ET,AL.

-""

I

---*--NH2- P. putida -C-

PP0301[PRO1031

NH4+

....... 0..... pH

E

?

100

lo% 7 .5 5

.............. 2.5

0

1 00 '

20

30 3 0

40

50

Days

lo

3i

7.5 5

+..................."', ..............-6

2.5

0

10

20

Days

30

40

1

FIG. 30. Kinetics of nitrogen transformation in Kitchawan soil amended with 6% montmorillonitg and inoculated with Pseudornonas putida PP0301(pR0103) or the homologous plasmidless parent P. putida PP0301 (Joneset al., 1991).

83


---*--N H ~ - E. -----t

0

I

108[R388]

... .I

.......*"'..

I I

C/OaCae

NH4+

N03-

PH

1

lo

z

I

*...*

5

7.5

*..................

5

........-*

2.5

0 10

0

NH2--W

20

Days

30

,

40

50

E. cloacae 107

NH4+

I N03-

lo

5

7.5 5

*..................t.................'.

c.... ..............

2.5

0

10

20

Days

30

40

50

FIG. 31. Kinetics of nitrogen transformation in Kitchawan soil amended with 6% montmorillonite and inoculated with Enterobacter cloacae 108(R388)or the homologous plasmidless parent E. cloacae 107 (Joneset al., 1991).

-

NH4+

NOS

7.5

5

2.5

Days

L .

NO3-

5

lo

7.5 5 2.5

0

10

30

20

40

50

40

50

Days S""

---*'-NHZII

-

--C-

NH4+

...... *..

NOZ-

.......

. ,d

0


NOS

FH

..................I

10

2o

Days

30

FIG.32. Kinetics of nitrogen transformation in Kitchawan soil not amended with montmorillonite and inoculated with Enterobacter cloacae 108(R388) or the homologous plasmidless parent E. cloacae 107, or not inoculated (Jones et al., 1991). The data in Figs. 32 through 34 are from experiments conducted concurrently. 84

-

-

.

N03-

E cn 1w a

lo

Z

%

7.5

....... ..................

5

~

2.5

0 10

0 *""

-E a

20

Days

40

30

50

1 2Vi -

---*--. NH2-

E. cloacae 107

NH4t

... 9......

N02-

--Q-

N03-

, .......

100

lo

*..+"'*.. ....... #y .....

z

I

!

&

7.5

.................................

.......t.""..............5..................i

5

I

2.5

0 10

0

20

40

30

50

Days

-

---*--.NH2-


NH4+

Q'.._.. N02N03-

c...................

~

.............A.

5 2.5

0

10

20

Days

30

40

50

FIG. 33. Kinetics of nitrogen transformation in Kitchawan soil amended with 9% inontmorilloniteand inoculated with Enterobacter cloacae 108(R388)or the homologous plasmidless parent E. cloacae 107, or not inoculated (Joneset al., 1991). 85

&

lo 7.5 5 2.5

0

'10

I

---*.--. NH2--C

NH4+

... 9'.....

N02-

--L.

N03-

20

Days

30

40

50

E. cloacae 107

2

4 loo

1 lo%

z

7.5

..............'..." .................

....v

c

5

2.5

0

0

10

NHZ-----t

NH4i

L .

N03-

20

Days

30

40

50


lo% 7.5 5 2.5

0

10

20

Days

30

40

>

FIG. 34. Kinetics of nitrogen qansformetion in Kitchawan soil amended with 12% montmorillonite and inoculated with Enterobacter cloacae lOB(R388) or the homologous plasmidless parent E. cloacae 107, or not inoculated (Joneset al., 1991).


87

creased, regardless of the microbes added. Whereas the differences in the rate of nitrification between the microbial treatments were small and probably not ecologically significant, the effects of clay concentration were both statistically and ecologically significant. These results indicated that although the GEMs evaluated had no significant and persistent effect on nitrification, the technique was sufficiently sensitive to detect changes in the rates of nitrification that resulted from changes in the clay concentration of the soil. Similar results have been reported by Kunc and Stotzky (1980)and Macura and Stotzky (1980). The population dynamics of bacteria involved in the transformations of fixed nitrogen were determined in the same soil from parallel respiration studies (i.e., from the master jars). The numbers of nitrifying, nitrate-reducing, and denitrifying bacteria did not exhibit any consistent differences over 35 days that could be attributed to the GEMs or their homologous plasmidless parents. The numbers of denitrifiers and nitrate reducers remained essentially constant during the first 14 days of the study and then increased by one order of magnitude in both the glucose-pulsed (after 1 4 days) and nonpulsed soils (Figs. 14 and 15). The numbers of both ammonium- and nitrite-oxidizing bacteria decreased slightly on days 8.and 14 and then increased to the initial levels in both the glucose-pulsed and nonpulsed soil (Figs. 16 and 17). G. SURVIVAL OF GENETICALLYENGINEERED MICROORGANISMS AND THEIRHOMOLOGOUS PARENTS IN SOIL The detection of the added GEMs and their homologous parents varied between experiments. For example, in one representative master jar study with the Kitchawan soil (Fig. 351, the numbers of GEMS and homologous parents detected decreased during the inoculation, but some of the GEMs and parents were not consistently detected on each sampling day, even though they were detected on subsequent sampling days. For example, E. cloacae 107 was not detected on day 3, but it was detected on all subsequent days: E. coli W3110(R702) was detected on days 0 , 3 , 7 , 2 8 , and 35 but not on days 14 and 21; and P. putida PP0301 was detected on all days except days 3, 21, and 28. Enterobacter cloacae 108(R388) was not detected after day 14, even though the parent was. The reasons for the inability to detect this GEM after day 14 are not known. In the master jar studies with the xeric Millican soil from Oregon, both the GEM [P. putida PP0301(pR0103)] and the homologous parent (P. putida PP0301) were detected throughout the experiment (Fig. 36). The numbers of the GEM and parent detected fluctuated and decreased

FIG. 35. Detection of genetically engineered microorganisms, Enterobacter cloacae 108(R388), Pseudomonas putida PP0301(pRO103),and Escherichia coli W3110(R702),and the homologous plasmidless parents, E. cloacae 107, P. putida PP0301, and E. coli W3110,on different days after inoculation into Kitchawan soil amended on day 0 with 1% (w/w)glucose and pulsed (P) with 1% glucose on day 14. See Fig. 9 for details.

89


a.

0,

0 7

0,

-0

IT

PP030 1 (PRO 103)

4 1

0

r

5

10

15

20

25

30

35

40

45

50

55

DAY FIG. 36. Detection of the genetically engineered microorganism,Pseudomonas putida PP0301(pR0103), and its homologous plasmidless parent, P. putida PP0301, on different days after inoculation into Millican soil. The soil was either unamended or amended on day 0 with 1% glucose (w/w), 500 ppm, 2,4-dichlorophenoxyacetate(2,4-D), or glucose plus 2,4-D.

90

G. STOTZKY ET AL.

by approximately two to three orders of magnitude during the 53-day incubation. Although not statistically significant, the numbers of both populations detected remained higher in soil amended with glucose. The GEM maintained the plasmid, pR0103, through the 53-day incubation, as shown by phenotypic expression, DNA analysis, and restriction patterns of endonuclease digests (Short et a]., 1991). IV. Discussion

These representative results demonstrate that the techniques used were sufficiently sensitive and reproducible to detect changes, when they occurred, in microbial populations and their processes in soil that resulted from the introduction of GEMs. The results also indicated that the introduction of GEMs into soil without the substrates on which the products of the novel genes function or the specific inhibitors to which they confer resistance is insufficient to evaluate adequately the potential ecological effects of the GEMs. For example, the reduction in CO, evolution and in the number of fungal propagules and the enhancement of dehydrogenase activity occurred only in soil amended with 2,4-D and inoculated with the GEM F. putida PP0301(pR0103).These effects would not have been detected if the soil had not been amended with the substrate that the GEM had been engineered to catabolize. These unanticipated effects were not predictable from the phenotype of the GEM. These results also emphasize that with the current limited amount of knowledge, the potential ecological effects of GEMs should be evaluated on a case-by-case basis, not only for the GEM involved but also for the soil into which the GEM is to be released. For example, the results observed with P. putida PP0301(pRO103) in the xeric Oregon soil were not observed when the GEM was introduced into an agricultural soil that contained an indigenous microbiota capable of mineralizing 2,4-D (Short et a]., 1990). The effects of the GEMs, added either alone or with the substrate on which the novel genes function, on microbial populations and processes, with the exception of CO, evolution, numbers of fungal propagules, and dehydrogenase activity in soil w e n d e d with 2,4-D and inoculated with P. putida PP0301(pRO103), were generally transient. Although some of these transient effects were statistically significant, it is doubtful that they were ecologically significant. Even those statistically significant effects that were relatively long-term (e.g., reductions in CO, evolution and numbers of fungal propagules) may not be ecologically significant.


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The lack of methodologies and theories with which to determine whether an effect is ecologically significant constitutes a major deficit in microbial ecology in general and, specifically, in risk assessment of the release of GEMs to the environment. The development of appropriate theories and methodologies with which to identify ecologically significant effects should have the highest priority in future studies in microbial ecology. The EcD concept (Babich and Stotzky, 1983, 1985; Babich et al.,1983)is a step in this direction, but it also does not define how much of a response to a dose of an environmental perturbant is ecologically significant. Monitoring the survival of GEMs and, especially, the transfer of their novel genes to indigenous bacteria in nonsterile soil is difficult, as many indigenous bacteria appear to be becoming increasingly more resistant to many of the antimicrobials (e.g., antibiotics and heavy metals) to which the novel genes in GEMs confer resistance. Inasmuch as the homologous parental strains do not contain the resistance markers present in their respective GEMs, monitoring the survival of the parents is even more difficult. Nevertheless, the fate of the GEMs, their novel genes, and the parents must be monitored, to be able to attribute any changes observed in microbial populations and microbe-mediated ecological processes to the novel genes in the GEMs or to their normal phenotype as expressed by their homologous parents. The inability to detect the added GEMs or their parents on each day of sampling during an extended incubation of soil in the laboratory (or in the field) does not imply that the GEMs or their parents did not survive. The inability to detect these populations may have been the result of (1) a “viable but nonculturable” phenomenon (see Stotzky et al., 1990);(2)the overgrowth of the presumably selective media on that day by indigenous bacteria that were resistant to the concentrations of antimicrobials used; (3)the amount of time available to detect these populations concomitant with measuring changes in other microbial populations and their processes; or (4) an actual decrease in these populations below the level of detection. For example, in experiments designed to evaluate primarily the survival of and gene transfer by some of the bacteria used in the studies in the Kitchawan soil, the GEMs and their homologous parents survived more than 30 days (i.e.,the duration of the studies), although their numbers decreased, and they transferred their novel genetic information (by conjugation or transduction) to appropriate recipients added to the soil (no evidence of transfer to indigenous bacteria was observed) (e.g., Devanas et al., 1986;Devanas and Stotzky, 1986, 1988;Stotzky, 1989;Stotzky et al., 1990;Zeph et al., 1988).Consequently, it can be assumed that any changes observed in

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microbial activity, species diversity, enzyme activity, and other parameters evaluated in Kitchawan soil inoculated with the GEMS were the result of the introduced GEMSand their novel genes. In the Millican soil from Oregon, wherein the survival and ecological effects of only one GEM and its homologous parent, both of which contained sufficient highly selective markers for their detection in nonsterile soil, were studied, the survival of the GEM and its parent was more easily and consistently detected and related to changes in microbial populations and their processes. Inasmuch as relating the presence of a GEM or its homologous parent (as a control) released to the environment to any changes in that environment (e.g., in microbial populations and microbe-mediated ecological processes) has numerous implications (e.g., scientific, environmental, assessment of risks, legal), methods (e.g., highly specific markers detectable against the background of indigenous microbes) must be developed for monitoring the fate of any introduced microbes and their novel genes in the environment. Although many assays of microbial populations and processes evaluated showed no detectable response to the presence of the GEMs, even when the substrate on which the novel genes of one GEM function was added, it would be premature to eliminate these from the battery of assays that was developed to detect the ecological impacts of GEMS in soil. An insufficient number of GEMs, especially those constructed from indigenous soil bacteria, has been evaluated, and most of the GEMs evaluated either contained innocuous novel genes (e.g., antibiotic- or heavy metal-resistance genes) or were evaluated in the absence of the specific inhibitor to which the genes confer resistance or of the substrate on which the products of the novel genes function. Consequently, the current battery of assays, with perhaps some additions (e.g., more enzyme activities), should be further evaluated with more realistic GEMs (e.g., those that contain catabolic genes, as well as genes for enhanced dinitrogen fixation and toxin production). The GEMS evaluated in these studies were used because they were readily available when the studies were initiated. Moreover, the primary purpose of these studies was to evaluate the applicability and sensitivity of the various assays. These studies constitute the first broadly based investigation of the effects of GEMS on microbial populations and microbe-mediated ecological processes in a natural habitat. Although there have been a few other studies on the effects of GEMS on such processes, these studies have been restricted to the evaluation of only one or a few ecological parameters. For example, Wang et al. (1989) reported a significant,


93

but transient, increase in CO, evolution from nonsterile soil amended with lignocellulose and inoculated with Streptomyces lividans TK23:3651(pSES), which contains a plasmid-borne gene that codes for lignin peroxidase. However, the enhanced production of lignin peroxidase by this GEM may have resulted from the protoplasting procedure used in Its construction, and, as the GEM was also genetically unstable, the enhanced CO, evolution could not be definitively attributed to the novel genes. Scanferlato et al. (1989) and Orvos et al. (1990) found no statistically significant differences in the numbers of indigenous bacteria in water-sediment and soil microcosms, respectively, inoculated with Erwinia carotovora L-864, a spontaneous rifampin-resistant mulant of E. carotovora L-833 that contains a fragment of plasmid DNA bearing resistance to kanamycin. Nodulation of soybean roots by a nonmotile mutant of Bradyrhizobium japonicum generated by Tn 7 mutagenesis, which was similar to the wild type in growth rate (but with a longer lag phase) in culture, soybean lectin-binding ability, flagellar morphology, and nodulating capacity, was significantly less than nodulation by the wild type (Liu et al., 1989). Considering the potential benefits that might be derived from the introduction into soil and other natural habitats of GEMs constructed to accomplish a specific task, it is surprising that more studies have not been conducted on the potential impacts, especially unanticipated ones, of GEMs on the structure and function of the habitats into which they will be introduced. These potential impacts are the major concern iibout the release of GEMs to the environment. The studies described herein have demonstrated the suitability and sensitivity of a battery of assays with which to evaluate such potential impacts and have shown ihat, in one case, unanticipated effects did occur when a GEM was itdded to soil amended with the specific substrate on which the products of the novel genes function. V. Summary

The potential benefits from the use of genetically engineered microorganisms in the alleviation of numerous problems in agriculture, pest control, bioremediation of toxic wastes, etc., that are amendable to biotechnology are essentially unlimited. However, there is justifiable concern, both scientific and with respect to public policy, about the potential effects of the release of GEMs to the environment on the structure, function, homeostasis, and health of the environment. A few studies have investigated the survival, colonization, and function of GEMs and 1 heir novel genes in some natural, including terrestrial, ecosystems, but

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there have been no substantive studies on the effects of GEMs on these ecosystems. Even if a GEM survives in the habitat into which it is introduced, does the job for which it was designed, and transfers its novel genetic information to indigenous microorganisms, there should be little cause for concern unless the novel genetic information results in some unanticipated and untoward impacts on the environment. This is the bottom-line concern about the release of GEMs to the environment. This article describes some methods with which to evaluate the effect of GEMs on microbial populations and processes in soil. These methods are based primarily on those developed for agricultural and ecotoxicological applications. The purpose of this article is not to present results on the effects of specific GEMs on specific microbial populations and processes but, rather, to summarize the concepts and methods that have been developed and tested and to indicate the applicability, sensitivity] and reproducibility of the methods. The methods include those for determining (1)the metabolic activity and carbon mineralization by the soil microbiota, as measured by the evolution of carbon dioxide; (2) the transformations of fixed nitrogen by perfusion techniques: (3) the fixation of atmospheric dinitrogen by the acetylene-reduction technique; (4) the species diversity of the microbiota, using selective and differential media: and (5)the activity of selected enzymes, such as acid and alkaline phosphatases (to provide a measure of the cycling of phosphorus), arylsulfatases (to provide a measure of the cycling of sulfur)] and dehydrogenases (to provide another measure of metabolic activity). The methods were evaluated both at New York University and at the Corvallis Environmental Research Laboratory of the U.S. Environmental Protection Agency. A local soil was used in each laboratory, but the same GEMs and methods were used in both laboratories. The results demonstrated that the methods were sufficiently sensitive and reproducible to detect changes, when they occurred, in microbial populations and their processes in soil that resulted from the introduction of GEMs. The results also indicated that the introduction of GEMs into soil without the substrates on which the enzymatic products of the novel genes function or without the specific inhibitors to which the products confer resistance is insufficient to evaluate adequately the potential ecological effects of GEMs. For example, a reduction in respiration and in the number of fungal propagules and an enhancement of dehydrogenase activity occurred only in soil amended with 2,4-dichlorophenoxyacetate(2,4-D) and inoculated with a GEM, Pseudomonas putida PP0301(pR0103), that contained novel genes for the partial degradation of 2,4-D. These unanticipated effects were not predictable from the phenotype of the GEM and would not have been de-


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tected if the soil had not been amended with the substrate (i-e., 2,4-D) that the GEM had been genetically engineered to catabolize. Many of the effects on microbial populations and processes that were observed were transient. Although some of the transient effects were statistically significant, it is doubtful that they were ecologically significant. The question of ecological significance was not directly addressed in these studies, as there are no theories and methodologies available to determine what constitutes an ecologically significant effect on microbial populations and processes in soil or other natural habitats. This lack of appropriate theories and methodologies constitutes a major deficit in microbial ecology in general and, specifically, in risk assessment of the release of GEMS to the environment. The development of such theories and methodologies must be of high priority. ACKNOWLEDGMENTS Although the research described in this article has been funded, in part, by U.S. Environmental Protection Agency Agreements CR812484, CR813431, and CR813650 to G. Stotzky and New York University, it has not been subjected to the Agency’s review and, therefore, does not necessarily reflect the views of the Agency, and no official endorsement should be inferred. Mention of trade names or commercial products does not constitute endorsement or recommendation for use. The assistance of Drs. K. A. Short and R. J. King in some of these studies and the suggestions of Drs. J. L. Armstrong, R. H. Olsen, and R. J. Seidler are gratefully acknowledged.

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Biochemical Engineering Aspects of Solid-state Fermentation M. V.RAMANA MURTHY,"N. G. KARANTH,*' AND K. S. M. S. RAGHAVA RAot *Fermentation Technology and Bioengineering Discipline tProcess Engineering and Plant Design Discipline Central Food Technological Research Institute Mysore-570013, India

I. Introduction 11. Mass Transfer in Solid-state Fermentation Systems A. Interparticle Mass Transfer B. Intraparticle Mass Transfer C. Oxygen Diffusion D. Degradation by Enzymes 111. Heat Transfer in Solid-state Fermentation Systems IV. Influence of Bioreactor Design on Mass Transfer V. Heat Dissipation in Solid-state Fermentation Bioreactors VI. Role of Water Activity VII. Important Physical Parameters in Solid-state Fermentation A. Nature of Substrate B. Available Surface Area C . Particle Size and Shape D. Effect of Mass and Thermal Diffusivities VIII. Mathematical Modeling in Solid-state Fermentation Systems A. Kinetics B. Concentration Gradients C. Temperature Gradients IX. Experimental Measurements A. Biomass Estimation B. Gaseous Concentrations and Temperature C. Effective Diffusivity of Mass and Heat X. Conclusions XI. Nomenclature References

I. Introduction

A glance at the history of fermentation science and technology indicates that solid-state fermentation (SSF) processes were almost completely ignored in Western countries after 1940 due to the rapid devel'Present address: Chemical Engineering Division, Indian Institute of Technology, Madras, India. 99 ADVANCES IN APPLIED MICROBIOLOGY,VOLUME 38 Copyright 0 1993 by Academic Press, Inc. All rights of reproduction in any form reserved.

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opment of submerged fermentation (SmF) (Lonsane et al., 1982,1985). This situation has changed in the last ten years, which have witnessed a resurgence of interest in SSF processes throughout the world (Ramesh and Lonsane, 1990; Steinkraus, 1984; Lonsane and Ramesh, 1990), owing to the high potential of SSF techniques (Lonsane and Karanth, 1991).In contrast, in oriental and Asian countries there has been extensive economic exploitation of SSF processes. The commercial application of SSF can be divided into two types: (1)socioeconomic applications such as composting of wastes, ensiling of grasses, and upgrading of lignocellulosic products or staple foods and (2)profit-economic applications such as production of enzymes, organic acids, and fermented foods (Mitchell and Lonsane, 1991). Solid-state fermentation involves the growth of microorganisms on moist solid substrate in the absence of free-flowing water. The necessary moisture in SSF exists in an absorbed or complexed form within the solid matrix, which is likely to be more advantageous because of the possible efficient oxygen transfer process. In SSF, the water content is quite low and the microorganism is almost in contact with gaseous oxygen in the air, unlike the case of SmF. The water activity in the substrate is also important. Solid-state fermentation does not refer to the fermentation of solid substrates in a liquid medium, nor does it refer to the fermentation of slurries. The major difference between SSF and SmF is that in the former the substrate is a moist solid, which is insoluble in water but not suspended in liquid (primarily water), whereas in the latter the substrates are solids dissolved or submerged in the liquid. The solid substrates act as a source of carbon, nitrogen, and minerals as well as growth factors, and they have a capacity to absorb water, which meets the vital requirement for water by the microorganism. SSF simulates the fermentation reactions that occur in nature, which include wood rotting, composting, and food spoilage by molds. The SSF process in the context of this article mainly refers to one that is conducted under controlled conditions and is useful in producing valuable products like enzymes or secondary metabolites (Ulmer et al., 1981; Hesseltine, 1977;Bailey and Ollis, 1977). In SSF reactions, the bacterial and yeast cultures grow by adhering to the surface of the solid substrate particle while the filamentous fungi are able to penetrate deep into the solid substrate particles for nutrient uptake (Moo-Young et al., 1983).The solid substrate thus also provides anchorage to microbial cells. As the microorganisms in SSF grow under conditions closer to their natural habitats, they may be more capable of producing certain enzymes and metabolites that usually will not be

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produced in SmF. For instance, mycotoxin was found to be produced on moist wheat grains, but it could hardly be produced in SmF (Butler, 1975).The oxygen requirement for growth and metabolism of the culture is derived largely from the gaseous state and, to a lesser extent, also from that present in dissolved form in the water associated with the solids. SSF thus involves three phases, gas, liquid, and solid, making the situation relatively more complex. Another major difference between SmF and SSF is that in SmF all the substrate is equally accessible to the organism as it is completely dissolved, while in SSF much of the substrate is not accessible initially. Further, as fermentation progresses the net amount of accessible substrate will always decrease in case of SmF, but it may decrease or increase or even remain constant at different stages of growth in case of SSF (Knapp and Howell, 1980). Gas transfer rates were found to be much higher in solid substrate fermentations than those obtained for submerged cultures in similar gas environments (Bajracharya, 1978).This was attributed to the high interfacial area-to-liquid volume ratios of semisolid substrates as compared with the ratios for gas transfer in bubble aeration of submerged cultures. A distinct advantage of SSF for fungal enzyme production, when compared with SmF, is observed in terms of enzyme productivity, product recovery, and fermentor volume (Schwartzberg, 1980). This is mainly due to higher enzyme concentrations in the liquid phase in SSF, which permits enzyme recovery at considerably lower energy inputs. The higher concentrations are attributed to higher oxygen transfer associated with the larger interfacial surface-to-liquid volume ratios resulting from the distribution of liquid film on the surface of the solid substrate (Mudgett, 1980). In SSF, studies on identifying and understanding the factors affecting the growth of microorganisms are relatively few, in comparison with SmF. There is a lacuna in the engineering design of SSF due to the difficulty in experimentally measuring the key process variables. In SSF, the amount of solids involved is very high and the medium is heterogeneous. This makes accurate measurement of parameters such as cell biomass level, nutrient concentration, pH, and temperature extremely difficult (Moo-Young et a]., 1983). Although considerable information is available on SSF, the engineering aspects, in particular mass and heat transfer effects in these systems, have received scant attention. The available literature on SSF (also called koji fermentation) is primarily qualitative, and little attention has been paid to kinetic studies. The main objective of this article is to highlight the paucity of

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information in this regard in literature, while reviewing the available information. For the effective design of bioreactors, it is very important to have mathematical models that will be useful in predicting the performance of the bioreactors beforehand. In the literature, there have been some attempts at modeling SSF reactions. However, these have been confined to the modeling of the kinetics of the reaction only and not the interaction of heat and mass transport with bioreactions. In this article, the importance of mathematical modeling involving the interaction of transport phenomena with biochemical reactions in SSF systems is highlighted. II. Mass Transfer in Solid-state Fermentation Systems

For the biochemical reaction to take place efficiently in a bioreactor, nutrients and other essential materials required for growth and maintenance must be available to the microorganism, which often involves their physical transport in the medium. To understand mass transfer in SSF, it is helpful to compare the situation in SmF, wherein the environment is homogeneous and the supply of oxygen to the microorganism is through the liquid phase only. Oxygen is bubbled into the fermentation medium, and agitation is provided to break the bubbles and distribute the oxygen throughout the system. The dissolved oxygen concentration is uniform in the bulk of the medium while concentration gradients exist in the liquid film around gas bubbles and microbial cells (Aiba et a]., 1980).Although the fermentation medium is a Newtonian fluid in the beginning, as the fermentation progresses it may tend to become non-Newtonian in nature. As a result, the increasing viscosity and inherent poor solubility of oxygen make oxygen transfer difficult in SmF, Oxygen transfer coefficients can be increased by agitation and to some extent by aeration. In contrast, the SSF system is heterogeneous, and oxygen transfer is limited by a liquid film on the substrate surface (Bajracharya and Mudgett, 1980;Mudgett and Bajracharya, 1979).As there is no free water, no bulk mixing can be provided in the liquid phase: therefore, the interfacial area and oxygen partial pressure become crucial factors for effective oxygen transfer (Mudgett, 1980). Further, the liquid film on the substrate surface, in which microorganism grows and product formation takes place, is relatively stagnant. At the growth regions, oxygen concentrations decrease due to uptake by the microorganism. The decrease in concentration occurs along the penetration depth, which is defined as the zone where active metabolic


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growth takes place, and may reach zero at a certain depth. The penetration depth can be increased by increasing oxygen transfer. It was observed by Mudgett and Bajracharya (1979) that high oxygen pressures helped in high oxygen transfer and stimulated amylase production. Oxygen transfer through the film is mainly due to dissolution and diffusion. The other possibility is that the microorganism may get oxygen directly from the gaseous atmosphere, which would be an ideal situation for the growth of the microorganism. Whatever the mode of oxygen transport, it was observed that the transfer rates in SSF are higher than those realized in SmF (Mudgett and Bajracharya, 1979; Mudgett, 1980). In some situations, like compost fermentation, the external oxygen concentration plays an important role. It was observed that as the external oxygen concentration increases, the rate of substrate decomposition increases but at the expense of compost uniformity. MASSTRANSFER A. INTERPARTICLE The transfer of oxygen from the void fraction within the solid phase to the growing microorganism is the interparticle mass transfer (MooYoung et d.,1983). The volume occupied by the air within the substrate gives the void fraction, which itself depends on the substrate characteristics and the moisture content. The moisture content should be optimal. If it is too high, the void space is filled with water and the air is driven out, which creates anaerobiosis. At the other extreme, if the moisture content is too low, the growth of microorganism will be hindered. Mixing and aeration are good means of achieving interparticle oxygen transfer, under the given conditions of void fraction and moisture content. However, at high values of the void fraction, mixing and aeration are not that critical as the voids contain enough oxygen to sustain the growth of the cells. Aerobic microbial growth requires oxygen to be present for oxidative phosphorylation to proceed. This oxygen comes from the surrounding atmosphere and diffuses into the pores of the substrate bed. It was observed in case of composting that, even when the pore size of the compost is increased, there was still an inadequate gas exchange, and high concentration gradients were observed. Thus, it was suggested that even if the pile is loosely packed, the diffusion of oxygen being an important factor, mixing and aeration have relevance (Finger et a]., 1970). Therefore, mixing and aeration at regular intervals will be useful in releasing the entrapped carbon dioxide and resupplying the void spaces with fresh air. In view of the heterogeneity of SSF, continuous or intermittent mix-

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ing and aeration are often practiced in order to prevent the exhaustion of oxygen in localized regions of the substrate mass. It was reported that moderate agitation and mixing of substrate particles enhanced secondary metabolite production (Hesseltine, 1972).This is believed to be due to particle separation effects that provide higher interfacial areas for oxygen transfer (Mudgett, 1980). Lonsane et al. (1985)have summarized the advantages of agitation. However, at times agitation may also cause some adverse effects as it may disrupt mycelial growth by breaking up actively growing cells (Takamine, 1914). B. INTRAPARTICLE

MASSTRANSFER

Intraparticle mass transfer refers to the transfer of nutrients and enzymes within the substrate solid mass (Moo-Young and Blanch, 1981). The main aspects that need to be considered here are the diffusion of oxygen into the substrate containing the biomass and the degradation of solid substrate by enzymes secreted by the growing microorganisms. In dealing with intraparticle mass transfer the effectiveness factor (E,) is one of the most useful concepts. It is defined as the ratio of the observed reaction rate (robs)to the rate in the absence of any substrate concentration gradients. This concept, which helps in quantifying the diffusional limitations in heterogeneous catalysis, is also applicable to SSF systems. An important parameter required for the evaluation of the effectiveness factor is the Thiele modulus (4), which is a measure of the rate at which the substrate is consumed in relation to the rate at which it is supplied by the diffusion process (Satterfield, 1970). By making use of this concept, a hypothesis was developed to evaluate intraparticle mass transfer limitation for the case of first-order reaction rate kinetics (Weisz and Prater, 1954). Bischoff (1967)has developed an extended hypothesis to assess the mass transfer limitations by defining a generalized form of the Thiele modulus valid for any reaction order and particle shape. This could be very useful since in most of the cases the rates of the biochemical reactions are highly nonlinear. Experimentation to check the probable application of these criteria to SSF would be of immense use, while development of more appropriate criteria suitable to SSF would be a valuable endeavor.

C. OXYGEN DIFFUSION Oxygen diffusion into mold pellets has been extensively studied in SmF. However, the intraparticle mass transfer in a mold pellet is differ-


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ent from that involved in SSF, where the microorganisms grow on and into the solid substrate particles. Even so, the analysis of oxygen diffusion in mold pellets will help in understanding the situation in SSF (Moo-Young et al., 1983).Recently, Mitchell et al. (1990a)studied the growth of Rhizopus oligosporus on model substrates in SSF. They showed that diffusive processes limit the rate of growth, especially within the substrate. This is because, to reach the interior, oxygen must pass through the actively respiring biomass at the substrate particle surface and then diffuse through the aqueous phase within the substratum. In the literature, information on oxygen transfer capabilities of fermentors involving complex heterogeneous three-phase systems such as SSF is sparse (Metz et al., 1979;Charles, 1978). This is partly due to the inadequacy of the existing technique for measuring the oxygen transfer coefficients (k,a) in these complex situations. Recently Andre et al. (1988)have suggested an improved method for the dynamic measurement of mass transfer coefficients for SSF systems.

D. DEGRADATION BY ENZYMES Diffusion of enzymes and substrate fragments is another important aspect of intraparticle mass transfer in SSF. For the most part, the substrate is water insoluble, whereas the organism can utilize only watersoluble substrate for growth (Suga et a]., 1975;Huang, 1975;Mandels et al., 1974). For this reason, the action of extracellular enzymes in degrading the solid substrate into soluble fragments is a very important step in SSF. If the mass transfer resistance is very high, this could even be the rate-controlling step. The diffusion of enzymes is facilitated by the open pore structure of the substrate, and the degradation can happen inside the substrate. In this case, the water-soluble fragments of the substrate will have to diffuse out of the solid matrix into the bulk region, where further enzymatic action will take place and metabolizable compounds are formed. However, when the porosity of the substrate is low, the major portion of the degradation will occur at the outer surface of the substrate (Knapp and Howell, 1980;Humphrey et al., 1977).In either mode of enzymatic action, the solid and polymeric material are modified so that they enter the cell and serve as carbon or energy sources. Thus, utilization of solid substrates by microorganisms is affected by many such factors that are relatively unimportant for the growth of microorganisms in SmF where the substrate is soluble and can penetrate the cell membrane. Mitchell et al. (1990)described the mode of growth of Rhizopus oli-

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gosporus on a model substrate. They hypothesized a series of steps including release of enzymes from the mycelium, enzyme diffusion, and the process of degrading the substrate. However, they indicated the need for more work to characterize adequately the growth on natural solid substrates. Their descriptive model does not identify the ratelimiting step in solid-state growth, but it highlights the importance of enzyme diffusion for substrate degradation. 111. Heat Transfer in Solid-state Fermentation Systems

During SSF, in general, a fairly large amount of heat is evolved, which is directly proportional to the metabolic activities of the microorganism (Chahal, 1983). In the initial stages of fermentation, the temperature and oxygen concentrations are the same at all the locations of the SSF bed. As the fermentation progresses, oxygen diffuses and undergoes bioreactions liberating heat, which is not easily dissipated due to the poor conductivity of the substrate. With the progress of the fermentation, shrinkage of the substrate bed occurs and porosity also decreases, further hampering the heat transfer. Under these circumstances, temperature gradients develop in the SSF bed. In the case of composting in heaps, the gradients will be much steeper as the heat transfer is much poorer, and temperatures can rise to as high as 70°C. The transfer of heat into or out of the SSF system is closely associated with the metabolic activity of the microorganism, as well as the aeration of the fermenting system. The temperature of the substrate is very critical in SSF. High temperatures affect spore germination, growth, product, formation, and sporulation (Moreira et al., 1981),whereas low temperatures are not favorable for growth of the microorganisms and for the other biochemical reactions. Unfortunately, few attempts have been made to provide special equipment in order to achieve good heat transfer in SSF. The low moisture content and poor conductivity of the substrate make it difficult to achieve good heat transfer in SSF. Significant temperature gradients are reported to exist even when small depths of the substrates are employed (Rathbun and Shuler, 1983); hence it is very difficult to control the temperature of the fermentors on a large scale. In fact, heat dissipation is one of the major drawbacks of SSF in comparison with conventional SmF, where good mixing provided for efficient dispersal of sparged oxygen also serves to give better temperature control. The conventional techniques and concepts used for temperature control in SmF are not easily adaptable to SSF. This makes temperature control in SSF all the more difficult. In the case of SSF, temperature

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control is primarily accomplished by adjusting the aeration rate. If the temperature is too low, then decreasing the aeration rate enables the temperature to rise due to the respiration of the microorganisms. However, enough care has to be taken in order to prevent the oxygen from falling below the critical level that would adversely affect metabolic activity of the cells. On the other hand, if the temperature of the substrate is high, increasing the aeration rate promotes cooling of the fermentation system as the heat will be taken away by the air leaving from the system. This, in turn, reduces the moisture content of the substrate, which is not favorable for the growth of the organism. To compensate for this, air that is partially saturated with moisture is used for aeration. Considering the interdependency of temperature and moisture content, it appears that air can be used effectively for temperature control, especially in case of static SSF reactors (Grajeck, 1988; Narahara et a]., 1984). The heat generated in the fermentation medium is directly related to the dry substrate matter utilized in the fermentation process: Eh

= gMd

(1)

where E h is the total heat produced in the bioreactor, g the heat generated per unit mass of dry matter utilized, and Md the amount of dry matter utilized. To maintain the thermal equilibrium, if the heat is to be removed from the system by aeration, the amount of air (L,) required can be calculated from the energy of air in relation to the heat produced in the fermentation; that is,

L, = Eh/(Hz - HI) (2) where HIand H,are the enthalpies of air at the inlet and outlet of the bioreactor, respectively. The dependency of the air enthalpy on the temperature and moisture content can be given as (Grajeck, 1988) H

=

1.006T

+ 1.86T + 2500X,

(3)

where 1.006 is the specific heat of air (kJkg K), T the temperature (in "C), 1.86 the specific heat of water vapor (kJ/kg K), X, (kg/kg) the maximum water content in the air at temperature T, and 2500 the latent heat of water (kJIkg).Thus, the inlet and outlet air enthalpies can be estimated. Temperature significantly affects the maximum water vapor content in the air and the air enthalpy, whereas it affects the water activity of the substrate to considerably lesser extent, in accordance to the Clausius-Clapeyron formula (Grajeck, 1988). Terui et al. (1957, 1958, 1959) reported a high heap aeration process using an incubator pro-

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vided with a cooling device in the interior. The substrate was a porous, solid medium such as bran and sawdust molasses. Saucedo-Casteneda et al. (1990)also used changing aeration rates and increasing water content for the purpose of temperature control. IV. Influence of Bioreactor Design on Mass Transfer

In spite of the recent surge of interest in SSF processes, its immediate and extensive industrial exploitation is not yet completely realized. One of the major reasons for this situation is the lack of information on efficient bioreactor design. A critical and exhaustive analysis of the information available in the literature regarding the different features of the bioreactor designs for SSF process and the criteria for selection of a particular design for a specific process indicates that the information available is too meager as compared to that on SmF techniques. On a large scale, SSF reactions are carried out mainly in three types of bioreactors, namely, the tray fermentor, the packed bed fermentor, and the rotating drum fermentor (Fig. 1).In a tray fermentor, the solid substrate is placed in trays, stacked one over another, in a controlled atmosphere room called a koji room. The length and breadth of the trays are much larger than the thickness, which in general is approximately about 5 cm (2 inches). While the top layer of the substrate bed is exposed to the gas phase, the bottom of the trays may be closed or perforated. Humid air is circulated in the koji room. Forced air circulation does not exist through the space between two successive trays or through the substrate bed itself, and oxygen transfer occurs primarily by diffusion. Hence, the porosity of the bed and the gap between the trays are critical.

FIG.1. Schematic diagram of different types of solid-statefermentors (Arima, 1964).


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At the start of the fermentation, the inoculum is uniformly mixed with the substrate, and the oxygen concentration is the same at all locations of the tray. As the fermentation progresses, oxygen diffuses and gets consumed in bioreactions, resulting in gradients of oxygen concentration. Simultaneously, carbon dioxide is liberated by the bioreactions. As there is no special attempt or provision available for the dissipation of carbon dioxide, its movement occurs largely by diffusion, which also affects the oxygen transport. Further, as fermentation progresses, the substrate shrinks because of mycelial growth, affecting the porosity, which, in turn, affects again the oxygen diffusion, making the gradients steeper. In some instances, SSF reactions are carried out in heaps, for example, in composting. Here, the fermenting system is turned at regular intervals to expose new surfaces to oxygen in the atmosphere (Finger et al., 1970). In the above-mentioned cases of SSF, there is no effective control over oxygen diffusion. This problem can be minimized in a packed bed bioreactor (Saucedo-Casteneda et al., 1990),where there is a forced convection of gases. The relative magnitude of the gradients could be much lower in packed beds due to forced convection caused by incoming air. Further, the carbon dioxide (liberated during the bioreactions) will be purged out, allowing its replacement by air. However, the reduction in bed porosity with progress of the fermentation still remains a problem. The third type of SSF bioreactor is the rotating drum fermentor. In these fermentors, the heterogeneity of the system can be reduced to a large extent as compared with static trays or packed bed fermentors, which helps in reducing the macrogradients. This type of bioreactor comprises a drum-shaped container mounted on rollers, which act both as support and as a rotating device. The rotating speed of the drum is usually low, 1-2 rotations per minute. Different types of drum fermentors have been reviewed in literature (Lonsane et al., 1985). Microbial growth in drum fermentors was reported to be rapid and uniform (Takamine, 1914;Underkofler et al., 1939;Schulza, 1962). However, sometimes the abrasion due to the tumbling of the substrate particles, encountered in this type of bioreactor, can break the mycelium, thereby hindering growth (Lonsane et al., 1985; Saucedo-Casteneda eta]., 1990). Hrubant et al. (1976)used a slow rotating, three-chambered drum fermentor containing baffles, which slowly push the fermenting substrate along the length of the fermentor. This arrangement facilitated the addition of substrate from the reservoir to the first compartment and removal of the fermentor product from the third compartment so that a continuous fermentation could be carried out. Laukevics et al. (1984)

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FIG.2. Rocking drum reactor for solid-state fermentation (Ryooet al., 1991).

have used different types of rotating drum fermentors and observed that properly aerated stationary layer SSF fermentors are simpler and perform better than those equipped with a mixer. They did not discuss the reasons for this; however, it could be due to the disruption of mycelia, which, in turn, causes a delay in growth. Recently, Ryoo et al. (1991)have developed a novel SSF bioreactor called the rocking drum reactor (RDR), equipped with an integrated computerized temperature-moisture control system. This is shown to overcome the drawbacks of growth inhibition due to mixing and mycelial disruption in conventional rotary fermentors and that of liquid percolation and clogging which reduces oxygen transfer in packed bed tower reactors or trickling filter reactors (Laukevics et al.,1984;Viesturs et a]., 1987;Barstow et al., 1988).In the RDR, a slow rocking motion is given to the reactor, during which substrate remains undisturbed but air and moisture are distributed evenly. An integrated control maintains constant temperature by blowing air through the substrate at constant velocity but varying relative humidity, forcing evaporation for cooling. Lost water is replaced by a cold water spray, regulated by a computer program based on the water balance equation of the system. A schematic diagram of the RDR is given in Fig. 2, and further details are described by Ryoo et al. (1991).They reported excellent control of the temperature and moisture content in the bioreactor. They further explained that the rocking motion helped produce an even distribution of the air and moisture in the substrate, overcoming


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the problems of air channeling and uneven drying in SSF reactors pointed out by other workers (Narahara et al., 1982;Tengerdy, 1985; Sat0 et al., 1983). Using the above RDR, Ryoo et al. (1991)were able to achieve higher biomass productivities and concluded that such an improvement could be even greater in large-scale SSF bioreactors. However, it needs to be pointed out here that RDR would perhaps involve more sophisticated reactor fabrication and control systems, and its ultimate economic feasibility needs to be examined and established. V. Heat Dissipation in Solid-state Fermentation Bioreactors

In general, several techniques can be adopted to strike a balance between temperature change and aeration rate depending on the design of the fermentor and the type of substrate. For instance, Silman et al. (1979)used a covered water bath, which is agitated by hand once a day for flask fermentations. In the case of column fermentors, temperature regulation was achieved by using a controlled temperature room or by circulating water in a jacket. In the case of bin fermentors, this was achieved by covering the roof with burlap that is continuously soaked with water. Nishio et al. (1979)have used a water bath in which a rotating drum type fermentor is steeped, and water, at constant temperature, is continuously sprinkled onto the fermentor. These methods have practical disadvantages. For example, the immersion water bath could be used only on a small scale. Constant temperature rooms may not be effective for large-scale fermentors because the thermal conductivity of the solid substrate is usually low, making the transfer of large amounts of heat energy generated and accumulated during the process of fermentation difficult. A general description of the tray fermentor and its operation is provided in the previous section. As the bioreaction progresses, a good amount of heat is liberated. Since there is no special attempt or provision available for the dissipation of this heat, the main mechanism of heat removal could be by conduction through tray walls (bottom and sides) and via the latent heat of vaporization of the moisture. Heat dissipation by natural convection is also a factor. In our opinion, considerable latitude exists for creating a better heat (and mass) transfer facility through a more efficient design of the tray bioreactor system, enabling, perhaps, forced aeration in the spaces between the trays as well as through the substrate bed in the trays. In the case of tray fermentors there is little control over temperature. As a result, considerable temperature gradients exist in the bed. The

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situation may be improved to some extent in packed beds, where there is forced convection of air. This is also a batch reactor, and the conditions at each point vary with time. It should be noted that continuous culture in SSF is difficult to implement with the present state of knowledge. The forced convection, mentioned above, reduces the temperature gradients to a large extent as the carbon dioxide is purged out by incoming air and the heat is carried away. Laukevics et al. (1984)have suggested different modes of heat removal in various types of rotating drum fermentors. For instance, an internal diffuser (circular chamber), apart from an outside jacket, can be used for more efficient heat removal. In some cases, a hollow shaft is provided at the axis of the fermentor through which cold water is circulated for heat removal. However, Laukevics et al. observed that heat removal was not an easily surmountable problem despite the internal and external cooling devices, leading to low productivity and high maintenance cost. The design of a bioreactor recently proposed by Ryoo et al. (1991), consisting of an integrated computerized temperature and moisture control system, which maintains a constant temperature by blowing air through the substrate at a constant velocity while varying its relative humidity, has already been discussed in the previous section. VI. Role of Water Activity

Water activity (a,) gives the amount of unbound water available in the immediate surroundings of the microorganism. While it is related to the water content of the substrate, it is not equal to the moisture content. Water activity is defined as the ratio of the equilibrium vapor pressure of the substrate (Pa)to that of pure water (Po) at the same temperature : a, = P,/PO (41 Water activity influences microbial growth and enzymatic and biochemical processes. It also affects microbial stability, as each organism has its own minimum water activity levels for metabolic activity. For example, the optimal water activity of fungi is about 0.7,yeast about 0.8, apd bacteria about 0.9 (Beuchat, 1981).A slight fluctuation around the optimal water activity value causes a large disturbance in the growth and metabolism of the microorganism. For the physiological activities of the microbes, the water activity of the medium rather than the moisture content is important (Gonzalez et al., 1988;Pirt, 1975). Water activity also indicates the water potential, which is the measure of the energy state of water, in the solid substrate. Water potential


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is of two types, osmotic and matric (Gervais et al., 1988). The former is due to dissolved solutes, while the latter is due to capillary forces. Thus, by monitoring the moisture content and the dissolved solute concentration, the water potential can be maintained at the desired level. Lindenfelser and Ciegler (1975) and Gonzalez et al. (1988) have shown, by investigating SSF of Aspergillus ochraceus, that, of all fermentation conditions, the initial moisture level is among the most critical. The benefit is twofold. First, the initial water content gives the water activity that is required for growth, and, second, it causes swelling of the substrate by which penetration by the mycelium becomes far easier for effective utilization of the substrate. However, the relationship between water content and productivity is not yet completely understood and needs detailed investigation. Although cell growth in SSF is considerably influenced by moisture, it appears that the water activity is a more fundamental parameter than moisture content for the growth of the microorganism. In this regard, Scott (1975) has shown the biological response to a particular water activity to be independent of the type of solute and the total water content of the substrate. It was observed that water activity decreases with fermentation time due to the evaporation of water on removing the metabolic heat of the substrate, formation of reducing sugars etc. These variations, perhaps, can be compensated for by the water activity estimation method proposed by Ross [1975) for a moist food, where the overall water activity is a product of the individual water activities of each ingredient, as follows:

. . (awn) a,, = (awl)(aw2)(aw3). (5) In the solid substrate the proportion of the bound and unbound water varies with temperature. Hence water activity, which is a measure of the water available in the immediate vicinity of the substrate, decreases with an increase in temperature. The reason for this could be that the solubility of solutes increases with the increase in temperature, thereby decreasing the available unbound water and in turn water activity. Silverman et al. (1983) showed that the growth rate of a particular bacterium is greater at lower water activity and higher temperature than that at higher water activity and lower temperature. However, they also observed that the limiting water activity for growth was lower at 37°C than that at 20°C. Large amounts of heat energy generated are accumulated in the fermentation medium due to the low thermal conductivity of the solid substrate. In practice, air is best suited for temperature and humidity control, which influences water activity, in SSF. Consequently, amounts

114


of cooling air in excess of that generally required for respiration of the microorganisms are to be employed in aeration. The amount of air required is estimated based on the value of the water activity to ensure optimal conditions for growth of microorganisms and metabolic heat production. At thermodynamic equilibrium, the relation between water activity of a medium and the relative humidity of air over the substrate is given by (Grajeck, 1988) a, = Ywa' =

m W m,

+ rn,

= -Pa =-

Po

ERH 100

where Y, is the activity coefficient, a' the molar fraction of solute, m, the number of moles of water, mi the total number of moles of all solutes, Pathe equilibrium vapor pressure of substrate, Pothe equilibrium vapor pressure of water, and ERH the equilibrium relative humidity. To maintain the water activity at the desired optimal level, the removal of moisture and heat, using air as the cooling medium, is critical. VII. Important Physical Parameters in Solid-state Fermentation

The physical factors that directly or indirectly influence microbial growth in SSF are the particle size, shape, surface-to-volume ratio, crystallinity, and porosity of the substrate. The physical morphology of the substrate, especially porosity and particle size, will govern the accessible surface area to both organism and enzyme (Knapp and Howell, 1980). The proximity of the organism is also important. If the organism is in the vicinity of the point of attack, especially when the organism is adsorbed on the surface of the substrate, then the transport path for the breakdown products remains short. The products tend to accumulate at the surface, increasing their local concentration at the surface. The growth will be fast until repression or inhibition by the products starts. In some instances, the organisms grow as a layer over the solid substrate surface, and the depth will increase to such an extent that it sets up resistance to oxygen or nutrient penetration. A. NATUREOF SUBSTRATE The nature of the substrate affects the SSF process significantly. The substrates utilized in SSF are, in general, natural, water-insoluble cellulosic or starchy materials. Even though such substrate materials are natural habitats for microorganisms, the heterogeneous nature of the unrefined solid substrate could adversely influence the kinetics of the reactions involved in the fermentation process.


115

Of late, attempts have been made to grow filamentous fungi on solid, inert materials impregnated with nutritive solutions. Here, the substrate and support are separate, and the support absorbs the liquid medium, which is advantageous. For instance, it is possible to use a liquid medium containing monomeric carbohydrates readily utilizable by the microorganism. The degradation of the solid matrix during growth can be avoided, and stable geometric conditions are ensured. However, most of the solid substrates being used are polymeric in nature such as polysaccharides, proteins, lignins, and nucleic acids, with others including pectate, hemicellulose, alginates in seaweeds, and chitin. The cytoplasmic membrane, however, does not normally allow the entry of large polymeric molecules unless they are broken down externally (outside the cell) into diffusible subunits. However, large polymeric molecules are produced by microorganisms and transported from inside the cell to the outside of cytoplasmic membrane. Some evidence suggests that proteins cross the membrane in a linear, unfolded, or partly folded form and attain their full tertiary structure only after passing through the membrane (Lampman, 1978;Costerton et al., 1974).The enzymatic hydrolysis of the substrate, and thus microbial growth on the substrate, is greatly influenced by the physical factors mentioned above.

B. AVAILABLE SURFACE AREA In SSF, because the substrate is insoluble, the rate of hydrolysis is dependent to a large extent on the available surface area rather than weight, within limits (Stone et al., 1969).Any new surface area generated by grinding, cutting, etc., must be accessible to enzyme molecules. For many reasons, the area available for gases is not the same as that available for enzymes. Therefore, separate methods must be developed for estimating this accessible area (Stone et al., 1969;Tarkow and Feist, 1969).Again, it is interesting to note that all the accessible area of the substrate is not completely susceptible to attack. An example is cellulose hydrolysis, which may be due to both physical and chemical factors such as crystallinity and lignin content (Knapp and Howell, 1980). For an enzyme reaction to take place, direct physical contact between the enzyme and its substrate must occur, producing an enzyme-substrate complex that then breaks down into products of the reaction. Therefore, it is to be expected that the rate of reaction should be a function of the surface area of the cellulose. However, the increase in the rate of cellulose hydrolysis is much greater than the increase in surface area would suggest. The excess rate may be due to the greater susceptibility of the inner layers of the substrate. Under conditions of availability of

116

M. V.RAMANA MURTHY ET AL.

nutrients, it was observed that available surface area has a prominent effect on growth of the organism (Mitchell et al., 1988). C. PARTICLE SIZEAND SHAPE

Another important parameter influencing microbial growth on solid substrates is the particle size. Humphrey et al. (1977),by working with Thermoactinomyces spp. on three different particle sizes of avicel, found a slight increase in the rate of growth and cellulose utilization with a decrease in particle size. Avicel, which is a regenerated cellulose, is a porous and relatively homogeneous substrate. Therefore, there could not be much difference in the surface area among the three particle size fractions, and the slight increase in microbial growth was attributed to the increase in mass transfer within the particles. Moreover, the initial size of the avicel particles did not relate to the initial degradation rates, which were instead related to the inoculum size. Hence the spherical shape (assumed) may not be the size commensurate with the overall particle dimension (Knapp and Howell, 1980).Electron micrographs of avicel have indicated a rough surface and not necessarily a microporous one, whereas other substrates, like wood, were found to have an essentially porous structure. It was also found that some changes in this porous structure occur during degradation due to enlargement of pore size within the lumen, which results in an increase in superficial area (Stone et al., 1969;Bungay et al., 1969). Another way of addressing this problem is the formation of a homogeneous model system by using agar gelatin, natural polysaccharides, or synthetic polymers (Weiss, 1973).In this case, the substrate particles as well as active cells are well distributed and then immobilized in the gelled matrix of the system. However, this may not be feasible on a large scale for commercial purposes, for many reasons. Another alternative is incorporation of the fermentation medium into a noninteracting, inert structured carrier such as plastic rings, spheres, or vermiculite (Aidoo et al., 1982). Although these artificial systems can mimic the real or true SSF system only to a certain extent, they may be explored as model systems in investigating the effects of many parameters.

D. EFFECTOF MASSAND THERMAL DIFFUSIVITIES In a homogeneous material containing two or more solutes whose concentrations vary from point to point, mass transfer takes place in the direction of decreasing concentration. The mass diffusivity or dif-


117

fusion coefficient (D) of a solute, which is the measure of its diffusive mobility, is defined as the ratio of its flux, J, to its concentration gradient as given in Fick’s first law, J =

-D- 6C 6Z

(7)

Fick’s second law, which is more frequently applicable to diffusion in solids than that in fluids, is more appropriate to explain mass transfer in SSF systems. It is given by the equation

where R, is the biochemical reaction term. However, diffusion in the solid matrix of the SSF substrate material is complex as it may actually be diffusion through the liquid/gas contained within the macro/micropores of the solid substrate. No information is available regarding the value of this diffusivity. It should be noted that Fick’s law is based on the assumption that the diffusivity is independent of concentration, which may not be true for highly concentrated systems. However, in SSF systems, the concentration of solute (oxygen)is usually low, hence it is still a safe assumption to say that diffusion of oxygen in SSF systems obeys Fick’s law, as also indicated by Georgiou and Shuler (1986). In SSF the diffusivity may not be constant as it strongly depends on the macro- and micropores of the substrate particle, which may vary during the course of fermentation. The interaction between the kinetics of the reaction [RT of Eq. ( 8 ) ]and the transport, through the parameter, diffusivity (D), needs to be known. Clearly, there is a need for systematic study of diffusivity in SSF systems, considering the above factors. Thermal diffusivity is a measure of how quickly the temperature will change when heat is generated in the solid substrate due to bioreactions. Materials with high thermal diffusivity will get heated quickly; thus, thermal diffusivity is an important property when considering unsteady state heat transfer situations in SSF systems. Several studies were made to understand the effects of physical parameters for efficient microbial growth in semisolid matrices (Finger et al., 1970).In a study made to check the effect of heat accumulation on the uniformity of a compost bed using insulated pads, this effect was shown to be minor, but the decomposition rate of the substrate was found to increase with temperature. However, when the external temperature was increased to about 320 K, there was a difference in the

118


uniformity of the compost, where the top part was decomposed at a faster rate than the lower part. Similarly, when the size of the compost pile was varied in the initial period of composting, large-size piles appeared to be more efficient. This is due to the increased temperatures, which, in turn, increase the reaction rates and oxygen transfer rates. However, the overall diffusion of oxygen per unit volume decreases with an increase in size. To account for these temperature variations while designing the bioreactor, the important physical parameter required is the thermal conductivity. This can be inferred from the value of thermal diffusivity of the substrate during fermentation. Several researchers have successfully developed techniques to determine the thermal diffusivity of foods, but few efforts have been made in the case of fermenting substrates. Lai et al. (1989)measured the thermal diffusivity of sorghum mash. In this case, it was found that the change in thermal diffusivity is not sensitive to the fermentation time. VIII. Mathematical Modeling in Solid-state Fermentation Systems

The overall bioreaction in SSF involves the transport of oxygen and carbon dioxide in the porous substrate on which the microbial biomass is growing. During the bioreaction, oxygen is consumed and carbon dioxide is produced, involving also the generation of heat. Thus, the reaction involves transport of oxygen into the interior of the substrate and transport of carbon dioxide and heat from the interior to the gas phase. Hence, in the reacting biomass, gradients of concentration and temperature build up. To achieve proper design of SSF bioreactors, with minimum gradients and high reaction rates, it will be necessary to have suitable mathematical models for the prediction of the course of the bioreaction as well as the gradients of concentration and temperature. Ngian et al. (1977)have shown the need for the elimination of mass transfer resistances in estimating the intrinsic kinetic parameters. Further, microbial growth is an overall effect of the interactions of the microorganisms and their environmental conditions. Hence, the laws of thermodynamics involving conservation of energy and mass are to be applied to such systems. Consequently, environmental conditions such as temperature, pH, osmotic pressure, concentration of products and nutrients, and oxygen levels affect the growth. Environmental control is relatively simple in SmF due to medium homogeneity, while in SSF it is much more difficult. As a consequence, serious problems of oxygen transfer, mixing, localization of pH, temperature, and nutrient levels occur. The important aspects of kinetics and interaction of mass and heat transfer with kinetics are highlighted in this article.


119

A. KINETICS Only a few reports are available regarding the kinetics of bioreactions in SSF systems. This is partly because of the technical difficulties involved in the measurement of growth parameters, analysis of cell mass, substrate consumed, product yields, etc., as well as physical analysis like growth patterns (Narahara et al., 1982). Some reports based on product formation kinetics (carbon dioxide evolution or enzyme secretion) are discussed below. 1. Kinetics Based on Carbon Dioxide Evolution

Sugama and Okazaki (1979) have obtained an expression for estimating the growth of Aspergillus oryzae cultured on solid media using an indirect method of determining carbon dioxide evolution during cultivation. For the logarthmic phase the following equations are given:

m = m, ept (10) where A is the amount (mg) of CO, evolved by respiration per gram of dry matter, a, the mg CO, evolved by endogenous respiratiodg dry matter, k, the mg CO, evolved by nonendogenous respiratiodmg dry mycelia formed, k, the mg CO, evolved by endogenous respiratiodmg dry mycelia/hour, m the mg dry mycelial weightlg dry matter, m, the initial value of m, and p the specific growth rate (h-l). On integration and for m values much larger than m, the following equations can be obtained: -A= k l + - k, m CL

By estimating the numerical values of the parameters in the above equations, the proportion of carbon dioxide derived from endogenous respiration in the logarithmic growth phase relative to the total carbon dioxide evolution (a,/A) was given as 0.098. To account for mycelial growth in the stationary phase (maximum value of m, denoted by N), Okazaki et al. (1980) introduced the logistic equation as an extension of their previous study. Thus, Eq. (9) becomes

_ dA dt

k1NE.L

1

+ ke-pt

1

k e-pt k2N + ke-pt + 1 + ke-pt

(13)

120


which on integration yields A =

klN 1 + ke-pt

k,N x +p

111-

ect + k k,N -i + k i + k

(14)

For sufficiently large values of t, Eq. (13) can be written as

k,=R 1 x -dA dt The parameters N and dAldt of Eq. (15) are estimated experimentally. In earlier work, Okazaki and Sugama (1979) found that the respiration quotient of the koji mold is nearly 1, using rice grains as the substrate, indicating that the oxygen consumption and carbon dioxide evolution are interconvertible. The experimental measurements of oxygen consumption were shown to be in good agreement with Eqs. (13) and (14). The observed oxygen consumption rate was not found to follow the same trend as that of glucosamine content (Aidoo et al., 1981), which is an indirect method for biomass estimation, or enzyme activity. This discrepancy was explained by stating that the oxygen consumption rate also contained the oxygen consumption rate for mycelial growth. Okazaki et al. (1980) have also shown the mathematical models to hold approximately in the case of wheat bran. Growth constants of Aspergillus niger were obtained for semisolid cultures on cassava flour by Carrizalez et al. (1981) in a packed bed microfermentor. The carbon dioxide produced was correlated to the specific rate of biomass growth. The following equation was written, considering that carbon dioxide was a product associated with growth, dP

V - = QpX, epi

dt where P is the CO, concentration, Qp the specific rate of formation of CO, ,X the biomass, t the time, and V the volume of sodium hydroxide solution in liters. Integrating Eq. (16), with the limits t = 0, P = pl0 yields


121

When P >> K', Eq. (18)reduces to

P = Kect Taking the logarithms of both sides of Eq. (19), we have log P = log(Q,X,/pV)

(19)

+ pt/2.3

(20) If the log of concentration of carbon dioxide versus time is plotted, a straight line is obtained, and the slope is pi2.3 (only the exponential phase is taken into consideration). 2. Kinetics Based on Enzyme Production

Recently, Mitchell et al. (1991a) have obtained an empirical model for growth of Rhizopus oligosporus on a model substrate in SSF. They proposed a direct relationship between enzyme activity and biomass production based on the stoichiometry of glucose conversion (assuming no accumulation of glucose within the substrate) as

dx- Y,E dt

where X is the biomass density (mg dry wt/cm2),t the time (hours), Y, the yield coefficient (mg dry wt/mg glucose), and E the enzyme activity (mg of glucose/hour/cm2). Experimentally observed enzyme activity was approximated empirically as E = r,t

Ol%) of various substrates resulted in a decrease in hydrogen evolution rates (Hirayama et al., 1986). No hydrogen production by immobilized cells of R. rubrum 7061 could be detected when the amount of malate provided to the nutrient medium was lower than 60 mM (Mignot et al., 1989~). The substrate concentration suitable for hydrogen photoproduction differed slightly according to the kind of substrate (Hirayama et al., 1986; Sasikala et al., 1991b). The appropriate concentration of various electron donors were as follows: malate, 0.4-0.5% (Hirayama et a]., 1986; Sasikala et al., 1991b);lactate, acetate, and butyrate, 0.2% (Hirayama et al., 1986);glucose, 0.5-0.7% (Margaritis and Vogrinetz, 1983;Singh et al., 1990);and alcohols, 0.02-0.1°h (Fujii et al., 1987). Miyake et al. (1982) observed hydrogen photoproduction from L-malate by R. rubrum at three different concentrations (25, 50,and 75 mM), and the rates decreased in the same manner regardless of the malate concentration; they concluded that the substrate concentration was not the factor limiting hydrogen photoevolution rates. Little is known about hydrogen evolution from mixed electron donors of known composition where these compounds are consumed simultaneously. Enhancement in photoevolution of hydrogen was observed with a combination of methanol + ethanol, or n-propanol + nbutanol (1:1 by volume) (Fujii et al., 1987), ethanol or n-propanol (16.6 mM) + malate (7.4 mM) (Fujii eta]., 1987),malate + propionate (1:3 by weight) (Vatsala, 1987a), butyrate (24 mM) acetate (60 mM) + lactate (40 mM) (Ma0 et al., 1986), butyrate (30 mM) + bicarbonate (10 mM) (Kim et al., 1981), succinate (14 mM) + Na,S (6 mM), and succinate (10 mM) + thiosulfate (10 mM) (Ohta and Mitsui, 1981).The rate of hydrogen evolution from mixed electron donors was higher than that from the individual compounds; furthermore, it was found that the total hydrogen evolved from mixed donors was more than the sum of the quantities from the individual substrates.

+

B. SUBSTRATE CONVERSION EFFICIENCY In addition to the rate of hydrogen photoevolution, the molar conversion of substrate to hydrogen is an important factor to be considered for practical exploitation of a particular anoxygenic phototrophic bacterial system, especially when pure chemicals are used for hydrogen photoproduction. Hydrogen production depends on the enzymatic activity of the substrate conversion for the electron supply. Theoretical maxima of

ANOXYGENIC PHOTOTROPHIC BACTERIA

225

hydrogen evolution from organic substrates are given by the following equation (Vincenzini et a]., 1981): C,HyO,

+ (ZX - Z) HzO =

x COz

+ ( y / 2 + 2~

- 2) Hz

The equations for various substrates studied are Acetate: C,H;O, + Butyrate: C4HnOz + Ethanol: C,H,O + Fumarate: c4&04 + Glucose: C$I,,08 + Glutamate: C,H,N04 + Lactate: c3&,03+ Malate: C4&,0, + n-Propanol: C,H,O + Pyruvate: C&ho, Succinate: C4H604

2 H,O

=

2 CO,

+ 4 H,

6 H,O = 4 CO,

+

3 H,O = 2 CO,

3 H,O = 4 CO,

+ 6 H, + 6 H, + 12 H, + 9 H, + NH3 + 6 H, + 6 H,

5 H,O = 3 CO,

+

4 H,O = 4 CO, 6 HzO = 6 CO,

6 H,O = 5 CO, 3 HzO = 3 CO,

+ 3 H,O + 4 H,O

10 H,

9 H,

=

3 CO,

+ 5 H,

=

4 CO,

+ 7 H,

The substrate conversion efficiency of anoxygenic phototrophic bacteria is expressed as a percentage of the theoretical maximum. The percentage conversion efficiency can be calculated from the substrate utilized and hydrogen evolved by comparing the hydrogen obtained to the theoretical maximal conversion as 100% using the following equation (Sasikala et al., 1990b): Oh

Substrate conversion efficiency = 100 x O/T

where 0 is the observed hydrogen production from substrate (mol) and T is the theoretical maximal hydrogen production (mol), calculated as s x n, where s is the substrate consumed (mol) and n the number of moles of hydrogen to be produced theoretically per mole of substrate used. Hydrogen production based on the amount of substrate supplied is called the “virtual yield,” and that based on the amount of substrate used is called the “real yield” (Stevens et al., 1986). Resting cell suspensions are more efficient converters of substrate to hydrogen, yielding efficiencies as high as 100o/o; growing cultures use part of the substrate for growth, resulting in a lesser conversion of substrate to hydrogen. Segers and Verstraete (1983) observed a conversion efficiency of 78% for R. vannielii metabolizing lactate during the first 10 days when growth was observed and 100% during the subsequent 10 days when the cells were in the nongrowing stage. However, growing cultures subjected to constant anaerobic conditions normally produce hydrogen at higher rates than those from washed cells (Weaver et a].,

TABLE V

Sussmm CONVERSION EFFICIENCY TO HYDROGEN PHOTOPRODUCTION FROM DIFFERENT SUBSTRATES BY ANOXYGENIC ~ O T O T R O P H I CBACTERIA Electron donor Acetate Butyrate Butyrate + lactate + acetate Ethanol Ethanol + malate D-Fructose D-GlUCOSe Gluconate DL-LaCtak?

DL-Malate n-Propanol n-Propanol Pyruvate Succinate Sucrose

+ malate

Organism

Conversion efficiency (%)

R. capsulatus R. capsulatus Rhodopseudornonasspp,

57-100 23-80 64-70

Stevens et d.(1983) Stevens et a]. (1983) Ma0 et al. (1986)

Rhodopseudomonas sp. 7 Rhodopseudornonas sp. 7 R. capsulatus Z1 R. sphaemides Glc + A. sphaemides Gnt + R. rubrum ATCC 11170 A. sphaemides

45 48 27 99 54-81 48-78

A . capsulatus

65-80

R. vannielii R. sphaeroides

78-100 57-100

R. capsulatus Z1 Rhodopseudomonas sp. 7 Rhodopseudornonassp. 7 R. palustris 42 OL R. capsulatus Z1 R. capsulatus Z1 A. capsulatus Z1

56 36 48 52 68 72

Fujii et al. (1987) Fujii et al. (1987) Hillmer and Gest (1977a) Macler et 01. (1979) Macler et 01. (1979) Segers and Verstraete (1983) Kim et al. (1982a, 1987a,b),Macler et a]. (1979) Hillmer and Gest (1977a),Willison et al. (1983). Stevens et al. (1983). Francou and Vignais (1984) Segers and Verstraete (1983) Macler eta]. (1979),Sasikala et al. (199Ob) Hillmer and Gest (1977a) Fujii et al. (1987) Fujii et al. (1987) Vincenzini et al. (1982a) Hillmer and Gest (1977a) Hillmer and Gest (1977a) Hillmer and Gest (1977a)

42

6

Reference


227

1980).A conversion efficiency of 0-100% from various electron donors to hydrogen has been observed among anoxygenic phototrophic bacteria (Table V). A recent report (Sasikala et al., 1990b) showed a conversion efficiency of more than 100% in R. sphaeroides, which was attributed to the simultaneous involvement of endogenous substrates; hence, it becomes essential to study the simultaneous hydrogen production from endogenous substrates for basic research on hydrogen metabolism as previously done by Gest et al. (1962). However, for studies on applied aspects, this may be negligible and hence may not be taken into account. Higher conversion efficiencies need not necessarily include higher hydrogen evolution rates (Stevens et al., 1983; Sasikala et a]., 1990b). Though a number of electron donors are used for hydrogen photoproduction studies, large-scale and outdoor experiments are almost entirely restricted to DL-lactate (Kim et al., 1982a, 1987a,b) because it is inexpensive and abundantly available and in addition generally produces high rates and conversion efficiencies. Stevens et al. (1983),who studied hydrogen evolution from DL-lactate, acetate, and butyrate by nine strains of R. capsulatus, observed that the average conversion efficiencies for the three H donors were, respectively, 55.5, 52.5, and 30.5%. In general, conversion efficiencies are high with organic acids, whereas carbohydrates are least efficiently converted to hydrogen. In comparison to organic acids, work on alcohols as electron donors is very scarce, but the conversion yields for mixed electron donors composed of alcohols and malate were higher than those of individual electron donors by Rhodopseudomonas sp. 7 (Fujii et al., 1987). A number of factors like gas phase composition (Sasikala et al., 1990b), stirring (Kim et al., 1981, 1987b), illumination intensity (Kim et al., 1982a; Stevens et a]., 1984) and duration (Kim et al., 1982b), temperature (Stevens et al., 1984; Fujii et al., 1987), pH (Stevens et al., 1986), altered carbon metabolism (Macler et al., 1979; Willison et al., 1984), loss of uptake hydrogenase (Odom and Wall, 1983), immobilization (von Felten et al., 1985), cell concentration (Vincenzini et a]., 1982a), nitrogen source used for growth (Ormerod et a]., 1961), and ammonia concentration (Odom and Wall, 1983) are reported to alter substrate conversion efficiencies. The gas phase of the assay was an important factor affecting the substrate conversion efficiency in R. sphaeroides O.U. 001 (Sasikala et al., 1990b), where the gas phases 1000/o H, and 10% N, in argon gas decreased the conversion efficiency from malate. In Rhodopseudomonas sp. (Kim et al., 1981) a conversion efficiency of lactate to hydrogen was calculated under three sets of conditions: (1)continuous illumination in stirred culture, 74.5%; (2) con-

228

K.SASIKALA ET AL.

tinuous illumination without stirring, 66%; and (3) periodic illumination (12 hour intervals) without stirring, 56.8%. Thus stirring increased the conversion efficiency, and periodic illumination diminished conversion of lactate to hydrogen. A similar effect of stirring was observed in R. sphaeroides B6 (Kim et al., 1987b), which produced hydrogen with a conversion efficiency of lactate to hydrogen of 73.9 and 61.6% in the cultures with and without stirring, respectively. Kim et al. (1982a) showed in R. sphaeroides B5 a conversion efficiency of lactate to hydrogen of 69.1% for a vertically placed reactor, which increased to 78% for an inclined reactor. The effect was attributed to the strong illumination and efficient agitation of the culture medium owing to evolved gases in the inclined reactor. Loss of uptake hydrogenase activity (Hup- mutants) was found to result in an increased stoichiometry of hydrogen evolution from glucose in R. capsulatus B10 (Odom and Wall, 1983), where 1.8 mol of hydrogen per mole of glucose was recorded for the wild type while for the Hup- mutant strain ST410 it was 2.4 mol. The difference was ascribed to the absence of hydrogen recycling in the Hup- mutant. In contrast, Willison et al. (1984), who observed a similar increased stoichiometry of hydrogen production from malate in a mutant strain (IR4) of R. capsulatus, concluded that this enhancement in conversion of electron donor to hydrogen is not due to lack of hydrogen recycling in the Hup- mutant (because, under the conditions employed, hydrogen recycling was not observed in the wild type), but rather is due to an altered carbon metabolism which affects the flow of reducing equivalents from organic substrates to nitrogenase. The level of enhancement in substrate conversion to hydrogen was dependent on the particular electron donor. The mutant strain IR4 produced 10-20% more hydrogen than did the wild type with DL-lactate or L-malate as major carbon sources, 20-50% more hydrogen with DL-malate, and up to 70% with D-malate. In an R. sphaeroides mutant, altered carbon metabolism was also found to be the reason for hydrogen formation in nearly stoichiometric amounts from glucose (Macler et al., 1979), whereas the wild type showed a conversion efficiency of only 24%. The wild type while metabolizing glucose accumulated gluconate, whereas the mutant strain did not appear to do so. Temperature enhanced the conversion efficiency of a mixed electron donor by Rhodopseudomonas sp. 7: with n-propanol malate at 30°C the conversion efficiency was only 48%, and at 40°C it was 70% (Fujii et al., 1987). In agar-immobilized R. palustris 42 OL, not only the maximal hydrogen evolution rate but also the efficiency of malate conversion decreased at higher cellular concentrations. From 62% at a cell concentra-

+


229

tion of 0.425 mg cell dry wt/cm3, the efficiency decreased to 58% at 6.0 mg cell dry wt/cm3 (Vincenzini et al., 1982a). Resting cell suspensions of sodium glutamate-grown R. rubrum showed a higher efficiency of malate conversion to hydrogen compared to ammonium chloride-grown cells (Ormerod et al., 1961). The stoichiometry of hydrogen produced to cellulose degraded in cocultures of Cellulomonas and R. capsulatus strain B l O O and its Hup- mutant strain ST 410 varied with ammonium concentration, the optimum being at 1and 2 mM, respectively (Odom and Wall, 1983). For R. sulfidophilus strain LMG 5202, the efficiency of malate conversion to hydrogen varied with the phosphate buffer concentration used, the highest being at 90 mM with the pH stabilized at 7.0 (Stevens et al., 1986).

C. OPTIMIZATION OF THE PROCESS Important factors regulating hydrogen photoproduction include pH, temperature, light intensity and wavelength, concentration of electron donor, age of the culture, cell density, and nutritional history of the cells. Hydrogen production can be studied as a two-step process (Hillmer and Gest, 1977b; Sasikala et al., 1991b): (1)a growth phase for the production of hydrogen-generating biomass and (2) a production phase from the biomass thus obtained. It can also be monitored as a single step, namely, simultaneous hydrogen photoproduction during growth (Kim et al., 1982a,b). The organic substrate used for growth has a clear influence on hydrogen photoproduction by resting cells from different electron donors (Fujii et al., 1987; Hillmer and Gest, 1977b; Vatsala, 1987a). Similarly, the mode of growth (lightjdark, aerobiclanaerobic) was also shown to influence the process. In general, high rates of production occurred when resting cells of dark aerobic grown R. sphaeroides O.U. 001 were used for hydrogen photoproduction (Sasikala, 1990). Mao et al. (1986), while studying the screening process for the isolation of photosynthetic bacteria with high rates of hydrogen photoproduction, observed that dark aerobic conditions were better than light anaerobic conditions. Cells of several anoxygenic phototrophic bacteria derived from colonies of aerobic cultures grown with (NH,),S04 in the dark produced hydrogen at higher rates than those grown with N, gas (anaerobic) under illumination (Ma0 et al., 1986). The nitrogen source used during growth is another important factor influencing hydrogen photoproduction by resting cells. Ammonium chloride-grown cells of R. rubrum produce hydrogen after varying lag periods, the length of which depended on the NH4+concentration re-

230

K.SASIKALA ET AL.

maining in the medium when harvested (Ormerod et al., 1961; Ormerod and Gest, 1962).For cells of R. rubrum harvested when approximately two-third of the ammonia initially present remained in the growth medium, and was used for hydrogen evolution from malate, hydrogen evolution began after a lag period of about 1 hour; thereafter, the rate slowly increased and became linear after 3-4 hours, with older samples showing progressively higher rates (Ormerod et al., 1961).The effect of a large number of amino acids on the photoproduction of hydrogen and growth revealed that L-aspartate, L-glutamate, L-proline, Lleucine, L-lysine, and L-arginine promoted growth and did not inhibit hydrogen photoproduction (Bregoff and Kamen, 1952);further, it was shown that DL mixtures of amino acids inhibit growth of anoxygenic phototrophic bacteria. Sodium glutamate used as a nitrogen source during growth proved to be more suitable for hydrogen photoproduction by resting cells of A. rubrum (Ormerod et al., 1961),with higher yields of hydrogen per mole of malate added and higher rates of hydrogen evolution than ammonium chloride-grown cells being reported. Similarly, enhanced hydrogen evolution was observed with dinitrogengrown cells of Chromatium sp. Miami PBS 1071 (Ohta and Mitsui, 1981)and A. rubrum (Ormerod et al., 1961).Periodic replenishment of the culture medium with yeast extract or trace elements from modified Hutner’s mineral base extended hydrogen production in R. sphaeroides up to 6 weeks, although the rate of gas evolution was reduced (Macler et al., 1979).The rate of hydrogen evolution was doubled or tripled by the addition of trace metals in Chromatium sp. Miami PBS 1071 with succinate plus thiosulfate as electron donors (Ohta and Mitsui, 1981; Ohta et al., 1981). Hydrogen photoproduction did not occur at pH values below 6.5 or above 8.0,and the optimal pH was about 6.8-7.5 for various strains of anoxygenic phototrophic bacteria (Ohta et al., 1981; Stevens et al., 1986;Margaritis and Vogrinetz, 1983;Willison et al., 1983;Peng et al., 1987;Sasikala et al., 1991b).The decrease in hydrogen photoproduction at a pH away from the optimum was much more pronounced at acidic than at alkaline pH. The final pH of the medium usually changes to higher values during hydrogen assays (Vincenzini et al., 1985;Stevens et al., 1986),which was explained as being due to consumption of substrate (an organic acid like lactate or malate) and to the low buffering power of the culture medium (Vincenzini et al., 1985;Stevens et al.,1986).The possibility of increased hydrogen production at alkaline pH (hydrogen evolution doubling on an increase of pH from 6.5 to 8.0) in R. capsulatus being due to increased nitrogenase activity was ruled


231

out because nitrogenase activity as measured by C,H, reduction showed only slight variations between pH 6.5 and 8.0 (Jouanneau et a]., 1980b). A high pH (8.5-9.0) is unfavorable for photoproduction of hydrogen because an active uptake hydrogenase functions optimally at this pH (Colbeau et al., 1978; Willison et al., 1983; Vincenzini et al., 1986; Peng et al., 1987). Continuous monitoring and adjusting of the pH to values around 7.0-7.5 or use of buffers was found to be essential for optimal hydrogen evolution. Increasing the phosphate buffer concentration resulted in an increased stabilizing effect of the final pH around 7.0, and the highest yield (33.6%) of DL-lactate conversion to hydrogen was reached with R. sulfidophilus strain LMG 5202 at a phosphate buffer concentration of 90 mM (Stevens et a]., 1986). Outdoor experiments with anoxygenic phototrophic bacteria are strongly affected by fluctuations in temperature and light intensity due to day-night cycles and seasonal, geographic, and climatic conditions. Stevens et al. (1984) studied the capability of different R. capsulatus strains for hydrogen production from acetate at different temperatures between 20 and 50°C and different light intensities between 100-15 W/ m2/second corresponding to midday (full sun) light and overcast daylight, respectively. They observed that the temperature and light intensity optima vary from strain to strain, and the selection of a particular strain must depend on the local climatic conditions. For outdoor cultures, an organism with a thermophilic or thermostable nature is desirable, especially in temperate and tropical regions for practical applications in order to minimize cooling of the cultures, which involves costly equipment and maintenance. For outdoor cultures, however, it is most desirable that the bacteria also be capable of producing good amounts of hydrogen at lower temperatures since it takes some time for the culture to be heated up; the rate of hydrogen evolution was very low in the mornings even though the light intensity was high (70 klux) for R. sphaeroides, for which the optimal temperature for hydrogen photoproduction was 40°C (Kim et al., 1982a). Kim et al. (1982b) selected R. sphaeroides B5, which showed the highest activity at 40°C and good activities at 20 and 30"C, for hydrogen production in outdoor cultures. In general, the temperature optimum for photoproduction of hydrogen is reported to be 30-35°C (Ohta et a]., 1981; Kim et al., 1982b; Zurrer and Bachofen, 1982; Sasikala et al., 1991b), and Watanabe et al. (1981) and Singh and Srivastava (1991) isolated thermotolerant strains of anoxygenic phototrophic bacteria which produce hydrogen even at elevated temperatures near 40°C. In anoxygenic phototrophic bacteria, light intensities higher than

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those supporting maximal hydrogen evolution do not result in a decline in hydrogen production, as observed among many cyanobacteria. The lowest light intensity supporting maximal hydrogen evolution, called the “light saturation point,” ranges from 6.5 to 20 klux for many strains (Table VI), though some can tolerate very high light intensities, the maximum recorded being 60 klux for resting cells of R. capsulatus (Willison et al., 1983). However, in R. rubrum a rapid increase in light intensity from low levels to saturating conditions caused a bleaching of the cells, and slow adaptation was needed (Zurrer and Bachofen, 1982). In R. sphaeroides (Macler et al., 1979) hydrogen production increased at an approximately linear rate in the range of 1000 to 12,000 lux, with the rate of production doubling with a doubling of illumination intensity, although intensities higher than 40 klux inhibited production of hydrogen gas. Rhodobacter sphaeroides 8703 evolved 151 pl/hour/mg dry wt at 10,000 lux (Ma0 et al., 1986);increasing the light intensity to 20,000 lux enhanced the rate to 262 pl/hour/mg dry wt (Miyake and Kawamura, 1987). Apart from an increase in the total amount and rate of hydrogen production with light intensity, the lag period also decreased for R. rubrum (Planchard et al., 1989). Hydrogen photoproduction by growing cells of R. capsulatus Z1 (Hillmer and Gest, 1977a) becomes “saturated” at an intensity of 6500 lux. An increase of light intensity during growth specifically leads to increased abilities in resting cell suspensions to produce hydrogen (Hillmer and Gest, 1977b), and hydrogen evolution in resting cell suspensions was saturated at TABLE VI LIGHTSATURATION FOR PHOTOPRODUCTION OF HYDROGEN BY ANOXYGENIC PHOTOTROPHIC BACTERIA

Light saturation Organism

(UUX)

Reference

R . rubrum A . rubrum S-1 A . capsulatus Z1 (resting cells) A . capsulatus Z1 (growing cells) R. capsulatus A. sphaeroides R. sphaeroides O.U. 001 R. sphaeroides 8703 R. rutila ATCC 33872 Chromatiurn sp.

15 12.245

Planchard et al. (1989) Zurrer and Bachofen (1982) Hillmer and Gest (1977b) Hillmer and Gest (1977a) Willison eta]. (1983) Macler et al. (1979) Sasikala et al. (1991b) Miyake and Kawamura (1987) Nogi et al. (1985) Mitsui (1981)

10.8 6.5 60

12 5 20

12 10.714


233

10,800 lux. The optimal light intensity for hydrogen production by R. capsulatus in continuous culture was much higher than that which saturated the growth energy demand (Vignais et al., 1984), whereas for batch cultures both hydrogen production and growth reached optima at the same light intensity (Hillmer and Gest, 1977a). The effect of light intensity on hydrogen evolution has been studied in R. rubrum (Stiffler and Gest, 1954), R. capsulatus (Hillmer and Gest, 1977b), R. sphaeroides (Sasikala et al., 1991b), R. rutila (Nogi et al., 1985), and Chromatium sp. (Ohta et al., 1981). In outdoor batch cultures of R. sphaeroides B5 illumination was found to be a limiting factor for hydrogen evolution (Kim et a]., 1982a). The culture, which was inclined at 30°, received more sunlight and produced more hydrogen with higher conversion efficiencies from lactate than cultures oriented vertically to the ground. Not only the intensity of light, but also its wavelength (Nogi et al., 1985) and source (Miyake and Kawamura, 1987) influence hydrogen photoproduction. A xenon lamp that caused a large increase in energy at 800-900 nm was found suitable for hydrogen photoproduction by Rhodobacter sp. 8703 (Miyake and Kawamura, 1987). Four major peaks of hydrogen evolution were observed at wavelengths of 900,860, 810, and 590 nm, and it was shown that wavelengths above 590 nm, which are absorbed by bacteriochlorophyll a, were more effective for hydrogen evolution than light of wavelengths below 540 nm, which are absorbed by carotenoids (Nogi et al., 1985). Hydrogen photoevolution in light-dark cycles of 16 hours light and 8 hours dark was unstable, so that hydrogen produced in the second light period reached only 50% of that produced in the first light period. This showed that uptake was highly active not only in the dark but also in successive light periods, causing a decrease of the net hydrogen productivity in the light, exceeding that occurring under continuous illumination. By removing hydrogen present in the reaction vessel with argon before the onset of the dark period, the rate of hydrogen production in successive light periods reached preceding levels (Vincenzini et al., 1986). Hydrogen evolution in three light-dark cycles (corresponding to 48 hours of light) was the same as that observed in 66 hours under continuous illumination owing to a greater hydrogen evolution rate; mean values in the light were 51 p1 hydrogen/cm2/hourin the light dark cycle and 35 p1 hydrogen/cm2/hourunder continuous illumination. The photosynthetic efficiency of hydrogen evolution by an anoxygenic phototrophic organism is the free energy of the total amount of hydrogen produced, divided by the total energy of the light incident on

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the cultures, times 100 (Vincenzini et al., 1982a; Miyake and Kawamura, 1987): Efficiency (Yo) = hydrogen output x hydrogen energy contentilight energy input x 100 Hydrogen production by photosynthetic bacteria requires organic compounds as electron donors whose free energy must be subtracted from the free energy of the hydrogen produced for an exact computation of energy balance (Vincenzini et al., 1982a). Photosynthetic efficiencies may be calculated as (Vincenzini et al., 1981) AGO'H,

-

AGO' SLE

in which AGO' H, and AGO' S are the free energies of the hydrogen formed and of the substrate converted, respectively, and LE is the light energy incident on the immobilized cells. The photosynthetic efficiency of R. palustris 42 OL at an incident light intensity of 69.5 call hour/100 cmzwas 1.2% (Vincenzini et al., 1981).This efficiency further varied with the cell density used. At 0.425 mg dry cells/cm3 the efficiency was 0.89% (average 0.62%), whereas at 1.7 mg dry wt/cm3it was 2.3% (average 1.6%). Relatively higher efficiencies were recorded for R. sphaeroides 8703 at higher light intensities (1000 W/mz), where maximum rates were observed. The efficiencies were 2.1 and 1.9% for a xenon and a filtered xenon lamp, respectively (Miyake and Kawamura, 1987). Efficiency became high at low light intensities; the maximum efficiency was 7.9% at 50 W/m2 using the xenon lamp and 6.2% at 75 W/mz using the filtered xenon lamp. The efficiency at low light intensities remained constant for long durations. At 75 W/m2via a xenon lamp, 99% of the initial value was maintained after continuous illumination for 10 hours, which is substantially as long as the duration of sunshine in a day. At high intensities degradation of efficiencies was observed; at 500 W/m2the efficiency decreased to 63% in 10 hours. Hydrogen evolution activity depends on the growth phase of the culture, and the highest activity was found for cells in midlog phase cultures (Ohta et al., 1981; Hirayama et al., 1986; Sasikala et al., 1991b). Increasing the concentrations of cells (0.2-1.6 mg dry wt/ml) increased hydrogen production (Sasikala et al., 1991b). The optimal cell concentration for hydrogen production was 1.6-1.8 mg dry wt/ml; higher cell densities resulted in lowering of the hydrogen evolution rate due to a self-shading effect. For agar-immobilized R. rubrum, a cell density of 1 mglml suspension before immobilization not only gives the highest short-term rates but, more importantly, much superior long-term activi-

ANOXYGENIC PHOTOTROPHICBACTERIA

235

ties (von Felten et a]., 1985). Marine Chromatium sp. Miami PBS 1071 was found to photoproduce hydrogen over a wide salinity range, with an optimum salinity of approximately 30% which is similar to seawater (Ohta et al., 1981). Elevated pressures of up to 2 bar were found to increase the hydrogen content of gases evolved by a culture of R. rubrum (Zurrer and Bachofen, 1982) because more carbon dioxide was absorbed by the medium. D. IMMOBILIZATIONTECHNOLOGY

A review of data available on immobilization of anoxygenic phototrophic bacteria for hydrogen photoproduction (Table VII) reveals that, although there is considerable disagreement regarding enhancement in the rates of hydrogen evolution, all reports point to the long-term stabilization of the process. Contradictory data are available on the enhancement in rates of hydrogen evolution by immobilization, with a 2to 10-fold increase in the hydrogen evolution rate reported, on one hand (Vincenzini et al., 1982a; von Felten et al., 1985; Ardelean et a]., 1989; Sasikala et al., 1990a; Singh et a]., 1990), and, on the other, an actual reduction in rates reported (Francou and Vignais, 1984; Matsunaga and Mitsui, 1982). Alginate-immobilized cells of Rhodopseudomonas sp. BHU 1 (Singh et al., 1990) and R. sphaeroides O.U. 001 (Sasikala et a]., 1990a) showed a 4-fold increase in hydrogen evolution rate over free cells, whereas a 2- to 10-fold increase was observed in agar (beads)-immobilized R. rubrum (von Felten et al., 1985). In contrast, carrageenan-entrapped cells of R. capsulatus B10 (Francou and Vignais, 1984) and agarimmobilized Rhodospeudomonas sp. Miami PBE 2271 (Matsunaga and Mitsui, 1982) retained only about 67 and 50% of the hydrogen evolution activity (rate) of free cells, respectively. In all the above cases, however, although the hydrogen evolution rate by free cells started to decline and completely ceased after some time, immobilized cells continued to photoproduce hydrogen at the same rates for a longer time. Continuous photoproduction of hydrogen was demonstrated for the following: carrageenan-entrapped cells of R. capsulatus at a rate of 3 ml/ hour for 16 days (Francou and Vignais, 1984), agar-entrapped cells of Rhodopseudomonas sp. Miami PBE 2271 at the same rate over 10 days (Matsunaga and Mitsui, 1982), agar-immobilized R. rubrum for 3000 hours with a loss of activity of 60% (von Felten et al., 1985), agarimmobilized Chromatium over a period of more than 300 hours (Ikemot0 and Mitsui, 1984), agar-cellulose fiber-immobilized R. rubrum

TABLE VII PHOTOPRODUCTION OF HYDROGEN BY IMMOBILIZED ANOXYGENIC PHOTOTROPHIC BACTERIA Hydrogen evolved (ccyhr/mg

Immobilization method Alginate

Agar beads Agar blocks

Agar sheet Carrageenan

Agarose coated polyester film with agar Agar cellulose fiber Agarose Pectin

Organism

Electron donor

dry wt)

Reference

R. sphaeroides O.U. 001 R. rubrum IF0 3986 R. rubrum Rhodopseudomonas sp BHU A. rubrum R. molischianum A. rubrum R. palustris R. sphaeroides 8703 R. palustris 42 OL R. rubrum 7061 R. capsulatus B10 R. rubrum G9 BM R. rubrum G9 BM R. rubrum Rhodopseudomonas sp. PBE 2271

Malate Acetate Lactate starch Lactate Wastewater Lactate Lactate Lactate Malate Malate Lactate Malate Acetate Lactate Malate

16.17 8.96 30.7 80

Sasikala et al. (1990a) Karube et al. (1984) von Felten et al. (1985) Sin& et al. (1990) von Felten et al. (1985) Vincenzini eta]. (1982b) von Felten et 01. (1985) Vincenzini et al. (1982b) Ma0 et al. (1986) Vincenzini et d.(1982a) Mignot et d.(1989~) Francou and Vignais (1984) Hirayama et al. (1986) Hirayama eta]. (1986) von Felten et al. (1985) Matsunaga and Mitsui (1982)

R. rubrum G9 BM A. rubrum A. rubrum

Acetate Lactate Lactate

57.3 139

5.5 50

151 41 0.23 111

9.35 18.04

29.3 445 15.6 22.9 21.0

Hirayama et al. (1986) von Felten et al. (1985) von Felten et 01. (1985)


237

B-9 BM for 60 days (Hirayama et al., 1986), and agar-immobilized R. palustris 42 OL for more than 60 hours at a rate only slightly lower than the initial one (Vincenzini et al., 1982a). Entrapment has been the method of choice for immobilizing anoxygenic phototrophic bacteria for hydrogen photoproduction. The advantages over adsorption are ease and reproducibility of immobilization as well as a high retentive capacity (Hallenbeck, 1983). Though a number of supports were tried for immobilizing anoxygenic phototrophic bacteria for photoproduction of hydrogen (von Felten et al., 1985; Hirayama et al., 1986), large-scale operations are almost entirely restricted to agar (Vincenzini et a]., 1986; Mitsui et al., 1985; Mignot et al., 1989a,b; Hirayama et al., 1986) and alginate to a lesser extent (Karube et a]., 1984; Sasikala et al., 1992). Von Felten et al. (1985), who screened different methods of immobilization, namely, agar (beads and blocks), agarose, potassium carrageenan, pectin, and calcium barium alginate for R. rubrum, observed that cells entrapped within small agar beads gave the best results when compared to others where the rates were about one-third to one-half those of agar beads. Cells entrapped in agar as a block were very ineffective, and calcium alginate as a support was not suitable for long-term experiments since no solid beads had been formed and the cells tended to wash out. Not only were the maximal and mean specific hydrogen production levels higher in agar, but the conversion ratio (hydrogen formed per lactate used) also increased significantly. In addition, the type of immobilization also affects the lag time for the start of hydrogen production, which was lowest for agar beads (von Felten et a]., 1985). Acrylamide gel (Weetall and Krampitz, 1980) and photo-cross-linkable resin (Hirayama et al., 1986), although stable, considerably iphibited the hydrogen-evolving activity of anoxygenic phototrophic bacteria. The geometry of the immobilized matrix is important and should maximize contact between the immobilized cell and substrate solutions so that limitations due to diffusion of substrates and products are significantly reduced; in other words, the matrix should have a high surface-to-volume ratio. Immobilization of R. rubrum in agar beads, which provides such a condition, gave by far the higher rates of hydrogen evolution with smaller lag periods when compared to agar blocks, due to the rather long distance for diffusion (von Felten et al., 1985).This may explain the reduction (Mitsui et al., 1985) and also lack of enhancement (Vincenzini et al., 1982a) in the rate of hydrogen evolution by immobilization in agar as slabs. The other advantage of beads is that they can be prepared more conveniently for use in large volumes and/ or in a fluidized bed reactor (Francou and Vignais, 1984). However,

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K. SASIKALA ET AL,

open-sided slabs were used successfully in reactors for various anoxygenic phototrophic bacteria (Vincenzini et al., 1981; Mitsui et al., 1985). Immobilized cells behave in a manner similar to free cells in terms of acetylene reduction (Hirayama et al., 1986; Sasikala et al., 1990a), effect of inhibitors and electron donors on nitrogenase activity (Sasikala et al., 1990a), and hydrogen production from a wide range of organic compounds in the light (Hirayama et al., 1986). Major factors specifically affecting hydrogen photoproduction by immobilized cells include the concentration of gel, diffusion of the substrate (and product), and light penetration, which are all interdependent apart from the factors affecting hydrogen evolution by free cells, though protection against various factors was observed with immobilization. Higher cell densities are known to cause substrate diffusion problems (Vincenzini et al., 1982a; Mignot et al., 1989a,b) and light limitation (Francou and Vignais, 1984). For agar-immobilized R. palustris (Vincenzini et al., 1982a), lower cellular densities of 0.425 mg dry wt/cm3 did not affect the substrate diffusion factor, and an average diffusion rate of approximately 32 p g malate/cm%our was observed. At densities higher than this, substrate diffusion became limiting, causing a diminution in hydrogen evolution. The possible limiting effect of light was ruled out because light saturated at 2.7 x lo3 erg/cm2/second for 0.425 mg cells/cm3 and much higher intensities were used. However, for carrageenan-entrapped R. capsulatus, the diffusional barriers appeared not to be rate limiting (Francou and Vignais, 1984); rather, there was light limitation. Light intensities that were optimum for free cells were found to be rate limiting for immobilized cells (Francou and Vignais, 1984) due to a shading effect resulting from the shadowing of beads by one another or from self-shading of bacteria inside the beads. At higher cell densities the latter becomes very important, causing limitation of hydrogen evolution. For agar-entrapped Rhodopseudomonas sp. Miami PBE 2271 (Matsunaga and Mitsui, 1982), limitation of light penetration into the gel was also thought to be the reason for hydrogen production rate limitation at cell concentrations higher than 800 pg protein/g gel at a light intensity of 150 microeinsteins/mz/second.For agar-immobilized R. rubrum (Karube et al., 1984), a cell content of 1% was found to be optimal. To trigger hydrogen photoproduction in R. rubrum immobilized in composite agar layer/micorporous membrane structures where the diffusional problem was very severe, up to 80 mM malate was necessary for noticeable hydrogen production to occur, whatever the light intensity (Planchard et a].. 1989). For R. palustris at a 0.8% gel concentration, substrate diffusion was not found to be limiting even at low


239

(20 mM) concentrations of rnalate (Vincenzini et al., 1982a), and the optimal concentration of substrate (acetate)for agar-immobilized Rhodospirillum sp. was 20 mM (Ardelean et al., 1989). Apart from substrate diffusion, the diffusion of product can also be a problem when immobilized living cells are used, where diffusion of gases rather than that of other substrates or products is the critical factor (Kierstan and Coughlan, 1985). For alginate-entrapped R. sphaeroides O.U. 001 (Sasikala et al., 1992), gas bubbles formed in the immobilized beads, as is usually observed when gas diffusion is slow compared to production (Krouwel and Kossen, 1980, 1981), which ultimately results in disruption of the gel matrix. However, no such effect was reported for beads of agar-cellulose fiber-immobilized R. rubrum G-9 BM (Hirayama et al., 1986) or carrageenan-immobilized R. capsulatus (Francou and Vignais, 1984) where continuous photoproduction of hydrogen was shown for 60 and 16 days, respectively. The concentration of gel to be used is another important factor when a balance has to be established between optimal hydrogen evolution (limited by substrate diffusion] and the mechanical strength of the immobilized matrix, which is very important for practical use. Thus, even though 1% alginate was optimum for hydrogen photoproduction, a 2% gel was used because the 1%gel was too soft for use with alginateimmobilized R. rubrum (Karube et al., 1984). With carrageenan as an immobilizing matrix for R. capsulatus (Francou and Vignais, 1984), however, no significant effect on hydrogen photoevolution was observed at the various carrageenan concentrations tested (2-4%, w/v). However, because making the beads with a 4% gel became difficult, due to the high viscosity of the liquid solution, and because at 2% the entrapment was not good, a concentration of 3.4% was selected. The concentration of gel is a very important factor affecting the substrate diffusion. The rate of hydrogen evolution in R. palustris was affected by gel concentrations; a 5-fold increase in hydrogen evolution was observed by changing the agar gel concentration from 3% (Vincenzini et al., 1982a) to 0.8% (Vincenzini et al., 1986) with other conditions substantially the same. Apart from enhancement and stabilization of hydrogen photoevolution, immobilization also enhances the storage stability of cells (Karube et al.. 1984; Matsunaga and Mitsui, 1982). Enhancement in hydrogen photoproduction rates on immobilization was found to be due to enhanced nitrogenase activity in alginateentrapped R. sphaeroides O.U. 001 [Sasikala et al., 1990a). Stabilization of hydrogenase and nitrogenase has been successfully achieved by immobilizing the heterotrophs Clostridium butyricum in polyacrylamide (Karube et al., 1976) and Escherichia coli in agar (Matsunaga et

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K. SASIKALA ET AL.

al., 1980) owing to protection of the hydrogenase and/or nitrogenase of the immobilized bacteria from the deleterious effects of oxygen because its diffusion was limited by the gel matrix. Photoproduction of hydrogen by immobilized cells of Rhodopseudomonas sp. PBE 2271 was markedly protected from the inhibitory effects of oxygen, nitrogen, and osmotic stress (Matsunaga and Mitsui, 1982). Stabilization of hydrogen evolution by immobilization of R. rubrum G-9 BM was thought to be due to protection of immobilized cells from oxygen and microorganic contamination and to the homogeneous distribution of cells in the gel (Hirayama et al., 19861. In addition, the pH of the medium surrounding the immobilized cells seemed to remain stable (von Felten et al., 1985). Although hydrogen photoevolution by immobilized anoxygenic phototrophic bacteria has been demonstrated over longer periods, a few problems are also encountered, including (1)deterioration of the gel structure with time (Hirayama et al., 1986), (2) cell leakage (Francou and Vignais, 1984;Planchard et al., 1989), (3) diffusional barriers (Vincenzini et al., 1982a; Planchard et al., 1989; Sasikala et al., 1992), and (4) light energy limitation (Francou and Vignais, 1984). To increase the strength of the immobilized cell matrix, various methods have been used, such as the inclusion of cellulose fibers in the agar gel matrix (Hirayama et al., 1986),immobilization of the agar matrix on an agarosecoated polyester film (Matsunaga and Mitsui, 1982), and applying a nylon net (pore size 50 pm) on the agar film with the purpose of sustaining the soft matrix (Vincenzini et al., 1986). To overcome the problem of leakage of cells into the medium observed with agar (Francou and Vignais, 1984), Planchard et al. (1989) have bounded the immobilized agar layer with a microporous membrane at the interface between the gel and the broth, which effectively checked the leakage problem and ensured confinement of cells in the gel matrix. However, the presence of the membrane decreased the rate of diffusion of the substrate from the medium into the gel; thus, a very high concentration of malate (80 mM) had to be used for hydrogen production. V. Enzymes Related to Hydrogen Metabolism

A. NITROGENASE Anoxygenic phototrophic bacteria have the unique ability of fixing dinitrogen, which is widespread among the various taxonomic groups (Madigan et al., 1984; Heda and Madigan, 1986a) with the exception of C. aurantiacus (Gogotov, 1985; Heda and Madigan, 1986b) and R. purpureus (Madigan et al., 1984). The enzyme complex nitrogenase cata-


241

lyzes the six-electron reduction of dinitrogen to ammonia with simultaneous reduction of protons to hydrogen: N,

+ 8H' + 8e- + 16ATP-,2NH3 + H, + 16ADP + 16Pi

Thus production of hydrogen is obligatory (Mortenson, 1978;Phillips, 1980),and though rates may vary it takes place at a minimal stoichiometry of 1 mol hydrogen evolved per mole of nitrogen fixed even under a nitrogen pressure of 50 atmospheres (Simpson and Burris, 1984). A minimum of 25% (possibly 40-60%) of the electron flux through nitrogenase during nitrogen fixation is directed toward production of hydrogen (Schubert and Evans, 1976);however, owing to the presence of uptake hydrogenase, which recycles hydrogen, production of net hydrogen may not be observed (Kelley et al., 1979;Zumft and Arp, 1981;Vignais et al., 1985).Unlike hydrogenase, nitrogenase produces molecular hydrogen irreversibly (Kosaric and Lyng, 1988).Hydrogen production represents a loss of energy (ATP) and reducing equivalents, and possession of an uptake hydrogenase seems energetically advantageous for nitrogen-fixing organisms (Dixon, 1972). However, in chemostat cultures of Rhizobium ORS 571, nitrogenasecatalyzed hydrogen production had more influence on the efficiency of nitrogen fixation than the absence or presence of a hydrogen uptake system (Stam et al., 1987).Nitrogenase also reduces other triple bond compounds including cyanide, acetylene, azide, hydrazine, alkyl isocyanides, and nitrous oxide (Burris, 1991).In the absence of any other substrate, nitrogenase reduces protons to hydrogen (Gottschalk, 1979; Bulen et QJ., 1965):

+ NADH + H + + H , + 5 A D P + 5 F / + NAD + 2e- + n m g A T P - H , + n m g A D P + 9

5ATP 2H+

Nitrogenase-mediated hydrogen production by resting cells of R. capsulatus BlO abruptly ceased on addition of C,H, (2.75mM), which was reversed by subsequent addition of CO. However, the C,H, reduction activity of the nitrogenase ceased completely on adding CO, indicating that CO is a selective inhibitor of C,H, reduction but allows proton reduction by nitrogenase (Jouanneau et QJ., 1980b);however, CO prevents hydrogen photoevolution by hydrogenase. In anoxygenic phototrophic bacteria most available evidence indicates that hydrogen photoproduction is mediated by nitrogenase: (1) hydrogen evolution like nitrogen fixation is light dependent (Gest and Kamen, 1949a;Kim et al., 1980); (2)hydrogen evolution had a clear correlation with nitrogenase activity (Watanabe et al., 1981;Miyake et

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al., 1982;Ohta and Mitsui, 1981;Macler et al., 1979)but not with hydrogenase activity (Kim et al., 1980);(3)hydrogen evolution is inhibited by dinitrogen (Gest and Kamen, 1949a;Macler et al., 1979),the natural substrate of nitrogenase; (4) hydrogen photoproduction was inhibited by acetylene (Macler et al., 1979;Ohta and Mitsui, 1981;Vincenzini et al., 1985),and the inhibition was removed by adding CO (Ohta and Mitsui, 1981,Jouanneau et al., 1980b); (5)hydrogen evolution is not inhibited by carbon monoxide, whereas classic hydrogenases are inhibited (Meyer et al., 1978a);(6)exposure to 18% oxygen in argon resulted in irreversible loss of both the hydrogen evolving and acetylene reduction capacities (Macler et al., 1979);(7)both hydrogen evolution (Macler et al., 1979) and nitrogenase activity (Kim et al., 1980)are similarly inhibited by ammonium salts, and the biosynthesis of the hydrogen-evolving enzyme is repressed in the presence of ammonia (Ormerod et al., 1961);(8) both nitrogenase activity and hydrogen photoevolution show an absolute requirement for ATP (Bulen et al., 1965); (9) like dinitrogen fixation, light-induced hydrogen evolution is suppressed in the presence of uncouplers and certain inhibitors of the electron transport chain (Kondratieva and Gogotov, 1981;Gogotov, 1985);(10)starvation of molybdenum results in a similar decrease of hydrogen production and nitrogenase activity (Kim et al., 1980);(11) the green filamentous thermophilic anoxygenic bacterium C. aurantiacus, which is incapable of nitrogen fixation, lacks the capacity for lightinduced hydrogen evolution as well (Gogotov, 1985);(12)nif- mutant strains which lack nitrogenase activity are incapable of hydrogen evolution although they contain an active hydrogenase (Wall et al., 1975; Siefert and Pfennig, 1978;Kim et al., 1980),and nif+ revertants capable of dinitrogen fixation also regained the ability to evolve hydrogen (Siefert and Pfennig, 1978);and (13)transfer of nitrogenase genes to nifstrains restored the capacity to fix N, and photoevolve H, (Solioz and Marrs, 1977;Wall et a]., 1975). Although nitrogenase activity is necessary for hydrogen evolution, the appearance of nitrogenase activity (in terms of C,H,) does not necessarily result in the simultaneous observation of hydrogen evolution (Peng et al., 1987). Simultaneous involvement of nitrogenase and hydrogenase in hydrogen photoevolution is observed under certain specific conditions depending on the growth phase (Vincenzini et al., 1985;Sasikala, 1990), the nitrogen source used for growth (Ohta and Mitsui, 1981),and regulatory mutants (Gorrell and Uffen, 1978);in the dark hydrogen evolution such simultaneous involvement is observed in light-grown cells (Voelskow and Schon, 1978).


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Nitrogenase (EC 1.18.2.1, reduced ferredoxin :dinitrogen oxidoreductase) consists of two oxygen-labile protein subcomponents, the molybdenum-iron protein (MoFe protein, dinitrogenase) and the iron protein [Fe protein, dinitrogenase reductase (Rc~)].Nitrogenase isolated from anoxygenic phototrophic bacteria (Ludden and Burris, 1976; Willison et al., 1983), are similar to those from other nitrogen fixers (Burgess, 1984) in size, structure including amino acid composition (Hallenbeck et al., 1982),and genetic material coding for nitrogenase (structural nif genes; Ruvkum and Ausubel, 1980). For nitrogenase activity (substrate reduction), the two nitrogenase proteins, ATP, a low-potential reductant, and an electron source are required (Stam et al., 1987; Zajic et al., 1978). The principal reductant (electron donor) in vivo is considered to be ferredoxin, the activity of which is the key function in the nitrogenase system for the photoproduction of hydrogen (Jee et al., 1987), and the electron source for ferredoxin reduction is either an organic substrate or inorganic sulfur compounds. Nitrogenase activity in anoxygenic phototrophic bacteria is strongly stimulated by light (Hillmer and Gest, 1977b; Jouanneau et al., 1985), although low levels of dark anaerobic (Madigan et a]., 1979) and dark microaerobic (Siefert and Pfennig, 1980; Madigan and Cox, 1982; Madigan et al., 1984) nitrogen fixation have been reported. The in vivo nitrogenase activity of R. sulfidophilus was maximal at a pH between 6.5 and 7.0 and decreased with increasing pH, resulting in reduced hydrogen production at pH 7.5 and none at pH 8.0 (Peng et al., 1987). In R. palustris, though nitrogenase activity was unaltered at higher pH, a reduction in hydrogen evolution was observed due to an enhanced uptake hydrogenase activity (Vincenzini et al., 1985). Nitrogenase synthesis in R. capsulatus is closely linked to incident light intensity (Vignais et al., 1984; Jouanneau et al., 1985). Unlike R. capsulatus (Meyer et al., 1978b) where resting cell suspensions prepared from aerobic dark-grown cultures showed nitrogenase synthesis during subsequent light anaerobic incubation, resting cell suspensions of aerobic dark-grown cultures of R. sulfidophilus failed to synthesize nitrogenase on light anaerobic incubation (Kelley et al., 1979).The control of nitrogenase synthesis by light might be energetic, the extent of nitrogenase derepression being a function of the energy available to the bacteria (Jouanneau et al., 1985). At light intensities higher than that sufficient to satisfy the energy requirement for optimal growth, a higher rate of nitrogenase synthesis was observed (Vignais et al., 1984). However, resting cells of R.capsulatus showed the highest stability of nitrogenase when they were exposed to a diurnal pattern of illumination

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rather than continuous light (Meyer et al., 1978b);they were capable of synthesizing the same level of nitrogenase as under continuous light, with a higher activity over a longer period. The ammonia fixed by nitrogenase from dinitrogen, or exogenously provided, is mainly assimilated through the glutamine synthetaselglutamate synthase (GS/GOGAT)pathway in anoxygenic phototrophic bacteria (Slater and Morris, 1974; Johansson and Gest, 1976; Bast, 1977; Weare and Shanmugam, 1976; Masters and Madigan, 1983; Herbert et al., 1978). However, R. purpureus, which is incapable of fixing dinitrogen, was found to employ the glutamate dehydrogenase (GDH)pathway as the primary means of assimilating ammonia under all growth conditions (Masters and Madigan, 1983). In anoxygenic phototrophic bacteria, nitrogenase activity is regulated in response to the availability of fixed nitrogen (Yoch, 1978) as also found in other diazotrophs (Laane et al., 1980; Cejudo et al., 1984; Kush et al., 1985; Yoch et al., 1988; Reich et al., 1986). When cells of anoxygenic phototrophic bacteria are exposed to ammonia, a short but quick inactivation of nitrogenase occurs that is short lived, sensitive, reversible, and has been termed the “NH4+switch off” (Zumft and Castillo, 1978). It is found in R. capsulatus (Hillmer and Gest, 1977b), R. sphaeroides (Jones and Monty, 1979; Yoch et al., 1988), R. rubrum (Schick, 1971), R. palustris (Zumft and Castillo, 1978), R. sulfidophilus (Kelley et al., 1979), A. viridis (Howard et al., 1983), and C. limicola f. sp. thiosulfatophilum (Keppen et al., 1985). Such rapid switch off was observed only in cells recently exposed to ammonia (Sweet and Burris, 1981); similarly, it is lost under N deficiency (Alef et al., 1981; Sweet and Burris, 1981). It was found to depend on the age of the culture (Alef et al., 1981; Sweet and Burris, 1981; Jouanneau et al., 1983). Growth conditions, such as light intensity (Yoch and Gotto, 1982), carbon source (Yoch and Cantu, 19801, and especially nitrogen source (Alef et al., 1981; Sweet and Burris, 1981);all regulate the rate and occurrence of the NH,+ switch off. Nitrogenase from anoxygenic phototrophic bacteria grown on dinitrogen or glutamate was isolated in an active or inactive form (Carithers et al., 1979; Ludden and Burris, 1976; Nordlund et al., 1978; Zumft and Castillo, 1978; Ludden et al., 1982a,b; Gotto and Yoch, 1985b; Yakunin and Gogotov, 1988). Although initially adenylylation/deadenylylation of glutamine synthetase was suggested to cause the nitrogenase switch off/on (Hillmer and Fahlbusch, 1979), Alef and co-workers (Alef et al., 1981) concluded that the two effects (ade/deadenylylation of GS and nitrogenase activity) are independent, even though both do usually respond to the same elicitor, namely, ammonia, as also confirmed by Yoch


245

and Cantu (1980). It was demonstrated in R. capsulatus (Jouanneau et al., 1983, 1989; Hallenbeck et al., 1982) and R. rubrum (Ludden and Burris, 1978, 1979; Gotto and Yoch, 1982b; Preston and Ludden, 1982; Ludden et al., 1982a, 1988; Pope et al., 1985; Lowrey et al., 1986) that inhibition of nitrogenase activity (switch off) had resulted from the inactivation by covalent modification of dinitrogenase reductase ( R c ~ ) , the Fe protein component of nitrogenase. The modifying group was found to consist of adenine, phosphate, and pentose (Ludden and Burris, 1978; Hallenbeck et al., 1982; Nordlund and Ludden, 1983). It was further shown to be an ADP-ribose molecule covalently linked to a specific arginyl residue of the Rc2 protein (Pope et al., 1985). It has recently been discovered in R. capsulatus that the specific residue of Rc2 involved is arginine 101, which is located in a highly conserved region of the polypeptide chain, and evidence for ADP-ribosylation of Rc2 was also presented (Jouanneau et al., 1989). This posttranslational ADPribosylation of Rc2 is carried out by dinitrogenase reductase ADPribosyltransferase (DRAT) (Lowrey and Ludden, 1988) in response to darkness or fixed N (Ludden et al., 1982a; Li et al., 1987) and a deficiency of organic compounds (Yoch and Cantu, 1980),thus inactivating the enzyme. Fitzmaurice et al. (1989) isolated genes coding for the reversible ADP-ribosylation system of Rc2 in R. rubrum. The inactive iron protein can be activated by the removal of the covalently bound modifying group by an activating enzyme, presumably an intracytoplasmic membrane-bound enzyme, in conjunction with a divalent metal ion and ATP (Ludden and Burris, 1976; Nordlund et al., 1977; Gotto and Yoch, 1982a, 1985a). The activating enzyme was purified and characterized as a specific arginine-(ADP-ribose) Nglycohydrolase (DRAG) (Saari et al., 1984; Pope et al., 1985). Liang et al. (1991) suggested the possibility of posttranslational regulation of DRAT and DRAG activities rather than transcriptional or translational control in R. rubrum. When ammonia is exhausted or light is again present, DRAG removes the ADP-ribose group and the nitrogenase is reactivated (Kanemoto and Ludden, 1984). Although it was suggested that in R. capsulatus the requirement for an activating enzyme was an artifact and that the activating enzyme played no role in the regulation of whole cell nitrogenase activity in anoxygenic phototrophic bacteria (Yakunin and Gogotov, 1983), Gotto and Yoch (1985a) have concluded that the activating enzyme does have a physiological significance in R. rubrum and that the phenomenon of activation of nitrogenase by the activating enzyme cannot be an artifact associated with the use of an artificial reductant. The alternative nitrogenase system in R. ru brum was also shown to be subject to similar regulation via posttranslational

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ADP-ribosylation of the Rc2 protein of nitrogenase (Lehman and Roberts, 1991),and the in vitro regulation of R. rubrum nitrogenase as well (Lowrey et al., 1986). The regulation of nitrogenase inactivation was found to be independent of the regulation of its synthesis in R. capsulatus (Hallenbeck, 1992).Yakunin and Gogotov (1988)assume that the formation of inactive nitrogenase is an initial stage of nitrogenase degradation under conditions of excess nitrogen. Apart from reversible ADP-ribosylation of Rc2, other mechanisms of nitrogenase inhibition have also been proposed (Lame et a]., 1980),including the inhibition of electron transport to nitrogenase (Haaker et al., 1982)and changes of the ATP/ADP ratios in the cells (Yoch and Gotto, 1982). Ammonia is also known to repress nitrogenase synthesis (Ormerod et al., 1961;Zumft and Castillo, 1978;Scolnick et al., 1983;Hoover and Ludden, 1984).Almost all the information hitherto indicates that nitrogenase regulation (its activity and synthesis) is directly related to the GWGOGAT ammonium assimilation pathway. In the presence of the glutamate analog L-methionine DL-sulfoximine (MSX), a powerful inhibitor of glutamine synthetase (GS), the inhibitory effect of ammonium on nitrogenase is alleviated in several strains of anoxygenic phototrophic bacteria (Weare and Shanmugam, 1976; Jones and Monty, 1979;Alef et al., 1981;Sweet and Burris, 1981;Zurrer eta]., 1981;Yoch and Gotto, 1982). Methionine sulfone, an inhibitor of glutamate synthase (GOGAT), potentiated the inhibition of nitrogenase by glutamine in R. sphaeroides (Jones and Monty, 1979),and 6-diazo-5-0x0-2norleucine (DON) an analog of glutamine which blocks GOGAT activity, was found to inhibit nitrogenase in R. rubrum (Yoch and Gotto, 1982).In the above cases glutamine accumulated due to inhibition of GOGAT and is thought to be responsible for nitrogenase inhibition. Thus L-glutamine has been considered as the metabolite responsible for regulation of nitrogen fixationhitrogenase function (Jones and Monty, 1979;Kim et al., 1980;Yoch and Gotto, 1982;Arp and Zumft, 1983; Michalski et al., 1983).Glutamine synthetase might play a role as an effector of the nitrogenase gene and regulate the level of nitrogenase activity in cells (Johansson and Gest, 1977;Wall and Gest, 1979;Hillmer and Fahlbusch, 1979;Yoch and Cantu, 1980;Alef et al., 1981;Falk et al., 1982;Engelhardt and Klemme, 1982;Jouanneau et al., 1984). Glutamine synthetase is thought to have a role because derepression of nitrogenase was observed either when GS synthesis was inhibited chemically by MSX (Meyer and Vignais, 1979;Moreno-Vivian et al., 1989)or genetically in GS-deficient mutants (Wall and Gest, 1979). It was shown in various anoxygenic phototrophic bacteria that the C/N ratio of the growth substrate influenced the derepression of nitro-


24 7

genase. Nitrogenase derepression, measured as hydrogen photoevolution, was observed in R. capsulatus Z1 at high ratios (1 or more) of external lactate to L-glutamate (Hillmer and Gest, 1977a) and as C,H, reduction in R. capsulatus ElFl (Moreno-Vivian et al., 1989) at high ratios (3 or more) of external DL-malate to L-glutamine. At a C/N ratio less than the above, free ammonia appears in the medium and nitrogenase is repressed. For agar-immobilized cells of R. rubrum, the maximal efficiency of hydrogen photoproduction was observed at a C/N ratio of 7 when m-malate and L-glutamate were the carbon and nitrogen sources, respectively (Planchard et al., 1989). Moreno-Vivian et al. (1989) concluded that nitrogenase synthesis in R. capsulatus E l F l is not regulated by ammonium per se, or L-glutamine alone, but rather by the intracellular C/N balance, and nitrogenase repression is triggered when the C/N internal ratio has shifted in favor of N, whose excess appears in the medium as ammonium. Nitrogenase is repressed (Hallenbeck et al., 1982) and inactivated (Jouanneau et al., 1980b) by oxygen. Both the MoFe and Fe proteins of nitrogenase are damaged by oxygen (Robson and Postgate, 1980). However, microaerobic and aerobic dinitrogen fixation is found in a few strains (Meyer et al., 1978c; Colbeau etal., 1980; Sasikala et al., 1991~). It was shown in R. capsulatus that oxygen may have a beneficial effect as an electron acceptor for the recycling of hydrogen evolved by the activity of nitrogenase, and 40% oxygen in the gas phase was required to inhibit in vitro nitrogenase activity completely (Meyer et al., 1 9 7 8 ~ ) . The mechanism of oxygen protection of nitrogenase is apparently mediation by respiratory activity (Hochman and Burris, 1981),and it has been shown in Azotobacter vinelandii to also involve catalase or another yet unknown factor(s) (Iwahashi et al., 1991). Nitrogenase activity is known to be enhanced in the presence of carbonate (Khanna et al., 1980);with alcohols as electron donors, bicarbonate or another suitable carbon dioxide source was found obligatory for nitrogenase activity in Rhodopseudomonas sp. 7 (Fujii et al., 1987). High concentrations of molecular hydrogen are known to inhibit N, fixation (Bothe and Eisbrenner, 1981), though low levels of hydrogen proved to serve as an additional electron source for nitrogenase activity (Meyer et al., 1978a; Sasikala etal., 1 9 9 1 ~ ) . The compact organization of nif genes found in Klebsiella is not observed in phototrophs (Haselkorn, 1986). For details regarding the genetics of nitrogenase, that is, the organization of genes for nitrogen fixation in anoxygenic phototrophic bacteria, see the review of Haselkorn (1986).Alternative nitrogenases (those with other metal Fe proteins in the place of the MoFe protein) have been found in Azotobacter vinelan-

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dii (Bishop et al., 1986;Hales et al., 1986;Robson, 1986)and the cyanobacterium Anabaena variabilis (Kentemich et al., 1988), both of which contain vanadium iron proteins and produce more hydrogen than the conventional enzyme complex. Searching for alternative nitrogenases in anoxygenic phototrophic bacteria is highly desirable and may enhance the value of solar energy conversion projects developed with them. However, substitution of tungsten (W) for molybdenum in R. rubrum resulted in a W-containing nitrogenase with lower hydrogen evolution than with the Mo-containing nitrogenase (Paschinger, 1974). Hydrogen evolution has not been studied in an alternative nitrogenase described by Lehman and Roberts (1991).An Fe-only alternative nitrogenase in R. capsulatus has been found by Schmider (1990). Continuous Synthesis of Nitrogenase For the practical exploitation of anoxygenic phototrophic bacteria for hydrogen production, stable and prolonged production of hydrogen is essential. Process optimization in terms of various environmental and physiological factors for hydrogen photoproduction alone is not enough to ensure prolonged hydrogen evolution at constant rates since the half-life of nitrogenase is rather low (40-50 hours; Bennett and Weetall, 1976;Kim et al., 1980;Moreno-Vivian et al., 1989). Continuous and constant synthesis of nitrogenase is an essential prerequisite for stable and sustained hydrogen photoevolution. Because ammonia and dinitrogen cause inhibition of hydrogen photoevolution by nitrogenase, these are usually omitted from hydrogen evolution experiments. As much as a 10-fold enhancement in nitrogenase activity was observed in nitrogen-deficient cultures of R. palustris as compared to N,-sparged cultures, owing to accumulation of nitrogenase rather than activation of the Fe protein by the activating enzyme (Zumft and Arp, 1981). However, in the long term nitrogen deficiency results in a reduced synthesis of nitrogenase, resulting in the loss of nitrogenase activity and hydrogen photoproduction. This can be overcome by additions of small amounts of nitrogen sources which do not inhibit hydrogen photoevolution by nitrogenase. Kim et al. (1980)recommended bubbling with nitrogen gas for the recovery of hydrogen photoproduction by a strain of R. palustris when hydrogen photoproduction had ceased in cultures. However, with R. rubrum (Miyake et al., 1982) it was found essential to add a nitrogen source [ammonium sulfate (0.1mM) or dinitrogen] when the culture showed a high activity of hydrogen evolution since the addition of nitrogen was not so effective after the bacteria had lost the ability for hydrogen production. Repeated addition of nitrogen sources could prolong the period of hydrogen evo-


249

lution under continuous illumination. However, because illumination is not continuous in the case of utilization of sunlight, the effect of nitrogen sources in the light and dark was studied, which revealed that supplying nitrogen at the beginning of the dark period is superior to that at the beginning of the light period (Miyake et al., 1982). In a simulated day-and-night system of la-hour light and 12-hour dark intervals, hydrogen production was extended and sustained for more than 2 weeks with periodic replenishment of nitrogen sources at the beginning of each dark period (Miyake et al., 1982). Similar work carried out by Vincenzini et al. (1986) for agarimmobilized cells of R. palustris revealed that the increased stability of hydrogen photoevolution allowed by light-dark cycles and by removing hydrogen from the photobioreactor before the onset of the dark period was further enhanced (stabilized] by removing hydrogen with dinitrogen in order to prevent nitrogen starvation of the entrapped cells. The best operational stability was achieved by addition of dinitrogen for 15 minutes in the photobioreactor at the beginning of each dark period and by substituting the substrate (20mM malic acid] every other day. This enhanced stability was also due to the maintenance of pH at neutrality, which favors hydrogen photoproduction (unlike a pH above 8) where uptake hydrogenase activity is high, though acetylene reduction does not change. For sustained photoproduction of hydrogen with growth, glutamate is known to be a good nitrogen source when used at concentrations ranging from 5 to 10 mM (Kim et a]., 1981, 1982a,b, 1987a,b; Mao et al., 1986; Miyake and Kawamura, 1987; Peng et al., 1987; Stevens et al., 1986). Another approach to maintain continuous production of hydrogen is to alter the ammonia assimilation system either chemically (by the use of MSX, an analog of glutamate; Weare and Shanmugam, 1976; Zurrer et al., 19811 or genetically (by nif-constitutive mutants; Weare, 1978; Liang et al., 19911, thus disrupting the tight coupling of nitrogen fixation and ammonia assimilation normally observed. B. HYDROGENASE

Apart from the participation of nitrogenase in hydrogen photoevolution, anoxygenic phototrophic bacteria also contain another hydrogenmetabolizing enzyme, “hydrogenase” (EC class 1.12), which catalyzes the reversible oxidation of molecular hydrogen. Determination of hydrogenase specific activity in the direction of hydrogen evolution is most commonly performed by following hydrogen gas formation in the presence of a suitable redox dye and an excess of dithionite to maintain

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a constant level of the reduced electron mediator. Another approach to measure evolved hydrogen is by a hydrogen electrode. Although most hydrogenases catalyze both hydrogen evolution and uptake depending on the experimental conditions, the reaction in the assay system containing dithionite and a suitable mediator (methyl viologen, benzyl viologen, ferredoxin, cytochromes, etc.) is considered unidirectional. It is also assumed that the initial rate of hydrogen production is proportional to the velocity of the enzyme at substrate saturation. Specific activity determined under these conditions should be independent of the enzyme concentration. It was found that the above fundamental postulates are not valid in the assay systems routinely used to determine the specific activity of hydrogenase. The apparent specific activity varies with the area of interface between the liquid and gas phase, and the apparent specific activity is strongly enzyme concentration dependent. A combination of these two effects interfering with the hydrogenase assay may bring about a 100-fold variation in measured specific activity (Bagyinka et al., 1984; Der et al., 1985). Because of the methodological flaws, an absolute comparison of the hydrogenase activities measured under various experimental conditions may be unintelligible. For example, specific activities determined for the same strain of T. roseopersicina in two different laboratories yield turnover times ranging from 6 to 2800 msecond (Der et al., 1985).Several discrepancies among data reported from various laboratories can derive from these effects. Hydrogenase activities have been detected in various compartments of the photosynthetic bacterial cell. In general, hydrogenases of photosynthetic bacteria are described as membrane-associated proteins. A good example of perplexing localization experiments is the C. vinosum enzyme. It has been described as strongly membrane bound (Feigenblum and Krasna, 1970;Gitlitz and Krasna, 1975;Kakuno et al., 1977),loosely membrane associated (Buchananand Bachofen, 1968;van Heerikhuizen et al., 1981),and soluble (Weaver et al., 1965).The ratio of “soluble” and “membrane bound” hydrogenase varies depending on the intensity of cell disruption (van Heerikhuizen et al., 1981).A similar behavior has been found in T. roseopersicina. Initially, two hydrogenases were reported for this bacterium, one soluble and one membrane bound (Serebryakova et al., 1977;Gogotov, 1978).A thorough characterization of the two enzymes revealed that they appear identical in all measured biochemical, physicochemical, and enzymological properties (Gogotov, 19781. In spite of the subsequent demonstration (Bagyinka et a]., 1982) of the existence of only one, membrane-associated hydrogenase, the confusion has persisted for some time (Gogotov, 1986). In R. ru-


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brum the situation is clear because the hydrogenase is so strongly embedded that one has to apply pancreatin treatment to remove the enzyme from the photosynthetic membrane (Gest, 1952;Adams and Hall, 1977, 1979). Yet, extracellular and soluble hydrogenase has been reported from this organism (Hiura et al., 1979).Rhodobacter capsulatus also contains a strongly membrane-associated hydrogenase which requires detergent treatment (Colbeau et al., 1978,1983)or extraction of lipids by acetone (Serebryakova et al., 1984) to be released into the aqueous phase. The localization of hydrogenase within the bacterial cell can be determined more precisely either by immunoelectron microscopy (Rohde et al., 1990;Lindblad and Sellstedt, 1990;Lunsdorf et al., 1991)or by redox dye permeability based methods (Jones et a]., 1976;Jones, 1980; Bagyinka et al., 1982;Kovacs et al., 1983;Kovacs and Bagyinka, 1990). The advantage of the first technique is the mild treatment conditions, whereas in the permeability based approach the orientation of functional active centers can be determined in addition to the compartmentalization of the enzyme. In a comparative study (Kovacs et al., 1983) the location and orientation of hydrogenase in the following photosynthetic bacteria have been established: T. roseopersicina, C. vinosum, E. shaposhnikovii, R. rubrum, R. capsulatus, R. viridis, C. limicola f. sp. thiosulfatophilum. Each of these strains displayed hydrogenase activity in both the hydrogen uptake and hydrogen evolution directions. Following a delicate enzymatic removal of the cell wall to form spheroplasts, and osmotic shock to disrupt spheroplasts, the hydrogenase activity sedimented with the membrane fraction in each photosynthetic bacterial strain except for C. limicola f. sp. thiosulfatophilum, where both the membrane fraction and the cytoplasmic supernatant contained hydrogenase activity. By applying a dye permeation method, samples containing benzyl viologen (BV) in the cytoplasm and cell suspensions where BV is accessible to the outer membrane surface only can be prepared. A simple measurement of the hydrogen uptake activity will then select the arrangement where hydrogenase and the redox dye BV are on the same side of the membrane. It has been found that in each species of purple bacteria, representing the various phylogenetic branches of photosynthetic prokaryotes, the hydrogen uptake center of the enzyme is located at the periplasmic side of the membrane. This spatial distribution of hydrogenase activity was later shown to support a bioenergetic advantage (Kovacs and Bagyinka, 1990). Chlorobium limicola f. sp. thiosulfatophillum may possess two hydrogenases. Alternatively, a portion of the same enzyme may be located in the membrane in the same orientation as in the other photosynthetic bacteria while another

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portion is loosely associated with the cytoplasmic side of the photosynthetic membrane. It is to be noted that an apparent discrepancy exists between these results and those based on immunological localization of the enzyme in R. capsulatus (Colbeau et al., 1983).A resolution of the conflict can be obtained by taking into account the fact that the two techniques look at differentparts of the molecule and thus the hydrogenase apoprotein may protude at the cytoplasmic side while the uptake center is embedded in the membrane and is accessible to redox dyes from the periplasmic side. Hydrogenases of several anoxygenic phototrophic bacteria have been purified and their catalytic properties studied. These include enzymes from R. capsulatus (Colbeau and Vignais, 1981; Colbeau et al., 1980, 1983; Seefeldt et al., 1987), T. roseopersicina (Gogotov, 1978; Gogotov et al., 1978; Kovacs et al., 1991a), C. vinosum (Gitlitz and Krasna, 1975; Kakuno et al., 1977; Llama et al., 1979; Strekas et al., 1980; van Heerikhuizen et al., 1981), and R. rubrum (Adams and Hall, 1977, 1979; Gogotov, 1978). Hydrogenase is supposed to contribute to the metabolism of anoxygenic phototrophic bacteria in the following ways. 1. Hydrogen recycling for enhanced efficiency of nitrogen fixation: As seen earlier, hydrogen evolution occurs obligately during dinitrogen fixation, amounting to a minimum of 25% (possibly 40-6096) of the electron flow through nitrogenase. The enzyme hydrogenase helps in recycling this hydrogen (Gogotov, 1978; Meyer et al., 1978a; Song et al., 1980), thus recovering some of the energy which would otherwise be lost (Dixon, 1972,1978). 2. Supporting autotrophic growth: Hydrogenase is necessary for the use of hydrogen as electron donor during photoautotrophic growth (Colbeau et al., 1978), and Hup- mutants lose the capacity (Vignais et al., 1984). During chemoautotrophic growth, hydrogenase enables the use of hydrogen as an electron donor and also an energy source with oxygen (Madigan and Gest, 1979; Siefert and Pfennig, 1979). 3. Scavenging oxygen and thus giving respiratory protection to nitrogenase: Meyer et al. (1978a) regarded energy and reductant generation as a minor function of recycled hydrogen compared with respiratory protection of nitrogenase. Vignais et al. (1984) have demonstrated that hydrogenase may be useful for hydrogen production by rapidly removing oxygen from the medium and thereby maintaining an anaerobic environment for nitrogenase, which is necessary for maximal activity of that enzyme. 4. Balancing energy requirements: Hydrogenase may contribute to


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the bioenergetic balance of the organism by generating membrane potential, namely, additional ATP from hydrogen (Kovacs and Bagyinka, 1990).

The in vivo uptake hydrogenase activity of R. sulfidophilus was pH dependent and at least partially responsible for the lack or low levels of hydrogen production below pH 6.75 and the results at pH 8.0 where the uptake was maximum (Peng et al., 1987). Hydrogen produced by the nitrogenase stimulates the synthesis of hydrogenase in growing cells of R. capsulatus (Colbeau et al., 1980). Unlike nitrogenase, hydrogenase was present in cultures grown on NH, +.It was demonstrated in R. acidophila that hydrogenase and nitrogenase are possibly linked genetically or by regulation (Siefert and Pfennig, 1978). Enzymatic properties can be best studied with purified hydrogenase protein. The results of protein biochemical studies usually facilitate genetic exploration at the DNA level. Interestingly, in very few cases are protein and DNA studies merged to understand the molecular mechanism of hydrogenase from a given photosynthetic bacterium. In Table VIII references to currently available hydrogenase molecular structure data are listed. It is interesting to note that the hydrogenases that are thoroughly characterized from a protein biochemistry and spectroscopy point of view (C. vinosum, T. roseopersicina) are not the primary targets of genetic analysis, whereas the proteins from the organisms containing known hydrogenase structural genes and/or genetics are poorly characterized biochemically (R. capsulatus, R. gelatinosus). Hydrogenases from various anoxygenic phototrophic bacteria show similarities in their subunit composition, metal centers, primary structure, and antigen determinants. One property shared by all hydrogenases is the possession of one or more iron-sulfur (FeS) centers as prosthetic groups. Most hydrogenases also contain Ni and a few accommodate Se in addition to the FeS and Ni centers. Those hydrogenases containing both Ni and FeS have an aP subunit structure, with the a subunit of molecular mass near 60 kDa and the /3 subunit of molecular mass near 30 kDa. Hydrogenases characterized from the anoxygenic phototrophic bacteria so far invariably belong to the group of a/3 NiFeS hydrogenases. Antibodies that specifically recognize hydrogenase protein structures have been prepared for a number of hydrogenases (Seefeldt et al., 1987; Kovacs et al., 1989). When the cross-reactivity of these antibodies was checked against various hydrogenases, strong cross-reactions were observed. These findings indicate a structural homology among the hydrogenases within the aP NiFeS group. In addition, structural similarities exist among a p NiFeS hydrogenases from

TABLE VID

STUDIESON HYDROGENASE MOLECULAR PROPERTIES AND CHARACTERIZATION FOR ENZYMESOBTAINED FROM P H o m s y ~ ~ ~ BACTERIA ~llc R. capsulatus Protein isolated Structural genes sequenced Genetic regulation of expression R. gelatinosus Structural genes sequenced R. rubnun Protein isolated C. vinosum Protein isolated Spectroscopic characterization (EF’R) T. roseopersicina Protein isolated Spectroscopic characterization

EPR PIXE EXAFS C. aumntiacus

Colbeau et al. (19831,Serebryakova et al. (19841,Seefeldt et al. (1987) Leclerc et 01. (1988),Richaud et al. (1990) Xu et al. (19891,Xu and Wall (19911,Toussaint et 01. (19911,Cauvin et al. (1991). Richaud et al. (1991) Uffen et al. (1990).Richaud et al. (1990) Adams and Hall (1977) Gitlitz and Krasna (19751,Kakuno et al. (1977),Strekas et al. (1980).van Heerikhuizen et 01. (1981).Serra et al. (1984) van der Zwaan Albracht et al. (1983,1984,1985,1986),Cammack et al. (1986a,b), et d.(1990) Gogotov et al. (19781,Kovacs et al. (1985,1988,1991a) Cammack et al. (1989) Bagyinlca et al. (1989) Maroney et al. (1990) Serebryakova et al. (1990)


255

anaerobic bacteria regardless of their taxonomic position or natural habitat (Arp et al., 1985;Kovacs et al., 1989). Additional strong evidence demonstrating the structural homology among ap NiFeS hydrogenases comes from their amino acid sequences (Leclerc et al., 1988;Uffen et al., 1990;Reeve and Beckler, 1990;Przybyla et al., 1992).Amino acid sequences deduced from the corresponding nucleotide sequences of the hydrogenase structural genes have been reported for R. capsulatus (Leclerc et a]., 1988) and R. gelatinosus (Uffen et al., 1990).The structural gene encoding the small (p) subunit is located upstream from the structural gene of the large (a)subunit as in the case of all known NiFeS hydrogenases (Reeve and Beckler, 1990). Moreover, the two hydrogenase sequences show extensive homology to each other and to sequenced membrane-bound hydrogenases from other bacteria. The amino acid sequences derived from the structural gene DNA sequences exhibit 70-75% identity. From an evolutionary viewpoint the most conserved parts of the FeS cluster-containing protein sequences are generally those involving the Cys residues that anchor the FeS clusters. Proteins that contain 4Fe4S clusters, such as hydrogenases (Beinert and Kennedy, 1989;Beinert, 1990),can often be easily recognized by the presence of a Cys-Xaa-XaaCys-Xaa-Xaa-Cys (Xaa = any amino acid residue) sequence which supplies three of the required four ligands to bind the FeS cluster (Bruschi and Guerlesquin, 1988).The fourth ligand is supplied by a single, conserved cysteine residue from some remote portion of the protein. As a result of this arrangement of ligands, FeS clusters generally bridge between distant protein segments. The genes for the large (a)and small (p) subunits of the sequenced NiFeS hydrogenases encode a variable number of cysteinyl residues, but none are organized in the -Cys-XaaXaa-Cys-Xaa-Xaa-Cys motif. There are, however, Cys-Xaa-Xaa-Cys motifs, and these are the ones now generally assumed to play a similar role in anchoring the FeS cluster(s) of the hydrogenase. Interestingly, these motifs are usually present at the amino and carboxyl ends of both subunits, and they appear to be highly conserved sequences. If they indeed turn out to be the ligands for the FeS cluster(s) one can expect a significant role of FeS clusters in stabilizing the three-dimensional protein structure of hydrogenases. There are at least two Cys-Xaa-Xaa-Cys sequences in all the large and small subunit sequences known so far, as well as a number of single conserved cysteines. The general tendency seems to be that more cysteines are located on the small subunit. Based on the relative abundance of conserved cysteines, it is often concluded that the FeS cluster(s) are likely to be bound to the small subunits (Reeve and Beckler, 1990).

256

K. SASIKALA ET AL.

In contrast to the above indirect evidence, direct measurements of the location of metal atoms on the individual subunits indicate that the Fe atoms are located on the large and the Ni atom is located on the small subunit of the T. roseopersicina hydrogenase (Bagyinka et al., 1989).A rational model incorporating both sets of findings delineates the metal centers in between the subunits, their binding involving residues from both subunits (Kovacs et a]., 1991b). At the present time, no biological Ni site has been crystallographically characterized. Several groups have reported the crystallization of NiFeS hydrogenases. However, no laboratory has yet reported any structural information from diffraction experiments. Questions to be addressed using the results of future crystallographic studies include the following. What is the exact location of the metal centers in the protein matrix? How does the absolute and relative arrangement of the redox centers change on transitions of the hydrogenase from one redox state to the other? Does the structural information on the interaction between the metals and protein provide clues that can be incorporated into the design of a functional model? Until direct answers to these and related questions are available, indirect methods, such as various spectroscopic techniques, must be applied to gain insight into the molecular mechanism of hydrogenase function. For this purpose, optical absorption spectroscopy is rather uninformative. Greater resolution is provided by electron paramagnetic resonance (EPR). The disadvantages of EPR are that it only detects the metal centers in those oxidation states in which they are paramagnetic, and it requires extremely low temperatures which preclude kinetic measurements. Initial hopes that one could detect the essential features common to all hydrogenases have been confounded. However, several different types of centers in hydrogenases can be distinguished. Based on'spectroscopic studies three sites or domains can be distinguished with recognizably different functions: the H-site, the A-site, and the R-site. Hydrogen activation takes place in the H-site domain. Because all hydrogenases use the substrate hydrogen, they are expected to contain one type, or at most few types, of such sites. Because the highly unusual metal Ni is found in all of these enzymes, it is a strong candidate for a component of the H-site. The first direct evidence indicating that hydrogen is in the coordination sphere of Ni was reported for the hydrogenase of C. vinosum (van der Zwaan et al., 1985). A light-sensitive nickel-hydrogen bond has been identified in this enzyme, which appears to be a general property of the NiFeS hydrogenases (Cammack et al., 1987; van der Zwaan et a]., 1987). Another common feature of hydrogenases is the inhibition of


257

enzyme activity by CO. Carbon monoxide changes the EPR signal of the hydrogen-reduced enzyme into a transient Ni-CO species. This Ni-CO species has been shown to be light sensitive, and photodissociation of CO results in the same nickel coordination as that after irradiation of the hydrogen-reduced enzyme (van der Zwaan et al., 1986). These results not only prove that hydrogen and the competitive inhibitor carbon monoxide bind to Ni at the same ligand position, they are the strongest indications that Ni is part of the hydrogen-activating site. Apart from identification of hydrogen and carbon monoxide as photodissociable ligands, there is more information on the coordination of Ni. Electron spin-echo envelope modulation measurements (e.g., Cammack et al., 1989, on T.roseopersicina hydrogenase) show only weak modulations of the Ni resonance due to 14N,probably resulting from a distal nitrogen of a histidine imidazole. The presence of sulfur in the direct coordination of Ni has been demonstrated by EPR and EXAFS (extended X-ray absorption fine structure) spectroscopy. The number of S donor ligands in the Ni coordination sphere is likely to be two (Maroney et al., 1990). The A-site portion of the enzyme harbors the electron acceptor. It is probably here that the different hydrogenases show the greatest diversity, because of the various electron acceptors that they use. The R-site serves as a regulatory site in the protein and accounts for the reversible inactivation of the hydrogenase under oxidizing conditions. To date the presumed regulatory site and the processes involved have not been directly identified. The extent, but not the first-order rate, of the activation of the oxygen-inactivated enzyme was found to be dependent on the redox potential of the incubation medium. Reduction of the Ni center proceeds relatively quickly, and hence it cannot be the rate-limiting step of the activation process; another redox center is assumed to function as a regulator site. The various relevant observations indicate that the regulation of activation involves a relatively rapid reduction of a redox center, followed by a slow conformational change in the protein which results in a change in the coordination state of Ni and allows the enzyme to react rapidly with hydrogen. Protein conformational rearrangements have been suggested to play a significant role in the redox regulation of the hydrogenases in photosynthetic (Tigyi et al., 1986; Kovacs et al., 1991a) as well as nonphotosynthetic NiFeS hydrogenases (Seefeldt and Arp, 1987; Moshiri and Maier, 1988). Most NiFeS hydrogenases are reversibly inactivated by oxygen, but some can become reactivated under reducing conditions. To describe the activation/deactivation phenomena three forms of hydrogenases have been proposed (Cammack et al.. 1986a,b).These forms of hydroge-

258

K. SASIKALA ET AL.

nases can be distinguished on the basis of their catalytic activities, spectroscopic characteristics, and protein conformational rearrangements (Kovacs et al., 1991a).The unready form is catalytically inactive toward hydrogen in all assays. Similar to this form, the ready form cannot catalyze the hydrogen isotope-exchange reactions either, but it can rapidly be converted to the active form by a strong reductant in the hydrogen-production and/or hydrogen-uptake assay mixtures. When assayed by hydrogen uptake, however, the ready form will not catalyze reduction of high-potential acceptors, such as dichlorophenolindophenol. The active form reacts with hydrogen without an induction period. The active form is produced rapidly by reduction of the ready form, but slowly by reduction of the unready form. EPR signals corresponding to the three functional states of the enzyme have been observed in a number of the NiFeS hydrogenases, which might imply that the activation/deactivation phenomena as described above are common to this class of enzymes (Fernandez et al., 1986; Cammack et al., 1989; van der Zwaan et al., 1990). Finally, a brief overview of the possible hydrogen activation mechanisms is presented in this chapter. Theoretically, there are three ways in which molecular hydrogen can be activated on interaction with a transition metal complex (M). Hydride formation Homolytic cleavage Heterolytic cleavage

-.

Mn+ H2 Mn+2(H-)2 2 Mn+ H, -. 2 MnflHM" + H, -. MnH- + H+

On activation of hydrogen, perturbation of the H-H bond can result in homolytic splitting of the hydrogen molecule into a pair of hydrogen atoms, each with one of the formal bonding electrons: H,+ZH-

Alternatively, H, can be cleaved heterolytically and, as both electrons are retained on one of the H atoms, a hydride and a proton remain: H,+H-

+ H'

Which of these mechanisms will occur largely depends on the oxidation state of the transition metal center and the nature of the ligands. Hydride formation, an example of concerted oxidation-addition, and homolytic cleavage both give rise to a formal oxidation of the metal center, and they require a low oxidation state of the metal and high basicity of the coordinating ligands. In the heterolytic activation mechanism, coordination of the hydride does not involve a change in the oxidation state of the metal, but the presence of an appropriate basic site


259

is required for stabilization of the release proton. This basic site may be external, as an added base, or the complex itself may contain such a site, leading to ligandlassisted heterolysis. Information on the mechanism of hydrogen activation by hydrogenase has been gathered through studies of isotopic exchange between D, and H+ and ortho-para hydrogen conversion (Krasna and Rittenberg, 1954; Teixeira et al., 1987). The observations can be brought in line with a heterolytic mechanism of hydrogen activation only if one assumes that the hydride and proton acceptor sites can exchange independently with the solvent. Hydrogenase enzymes isolated from various microorganisms show remarkable similarities in a wide range of structural properties. The comprehensive similarities, however, do not explain some pronounced variations, particularly in relation to enzyme stability. Elucidation of the structural basis for these properties is critical from a practical utilization point of view and requires further extensive studies. VI. Carbon Assimilation

Carbon assimilation by anoxygenic phototrophic bacteria depends on environmental metabolic variables like the substrate used for growth and/or hydrogen production (Hillmer and Gest, 1977a),aerobidanaerobic conditions (Schon and Voelskow, 1976; Schultz and Weaver, 1982; Harwood and Gibson, 1988), light/dark conditions (Kohlmiller and Gest, 1951; Glover et al., 1952; Clayton, 1955; Morita, 1961; Jungermann and Schon, 1974; Gorrell and Uffen, 1977, 1978; Voelskow and Schon, 19781, and the presence or absence and the specific nitrogen source used (Gest et al., 1962). Green sulfur bacteria are primarily autotrophic whereas purple nonsulfur bacteria are primarily heterotrophic (Ormerod and Sirevag, 1983), and the assimilation of CO, and organic compounds proceeds largely by different mechanisms in the two groups of organisms. Autotrophic assimilation of CO, in the light (photoautotrophic) and in the dark (chemoautotrophic) requires H, (or H,S) as the source of reducing power. In addition to the requirement of H, as a source of energy and reducing power, chemoautotrophic CO, assimilation requires 0, as the terminal electron acceptor for energy transduction (Madigan and Gest, 1979). Autotrophic CO, fixation by purple bacteria is carried out largely by the reductive pentose phosphate cycle (Fuller et al., 1961; Anderson and Fuller, 1967a; Ormerod and Sirevag, 1983), similar to that of higher plants and algae (Anderson and Fuller, 1967a). The occurrence of ribulose 1,5-bisphosphate carboxylase has been dem-

260

K. SASIKALA ET AL.

onstrated in both purple nonsulfur (Anderson and Fuller, 1967a,b; Hallenbeck et al., 1990a,b) and purple sulfur bacteria (Losada et al., 1960; Brown et al., 1981; Torres-Ruiz and McFadden, 1987; Heda and Madigan, 1988). In the green bacterium C. thiosulfatophilum, which lacks the key enzyme ribulose 1,5-bisphosphate carboxylase of the reductive pentose phosphate cycle, CO, is assimilated by the reductive carboxylic acid cycle (Evans et al., 1966; Buchanan et al., 1967, 1972; Sirevag and Ormerod, 1970a,b; Sirevag, 1974, 1975; Buchanan and Sirevag, 1976). Though there had been some controversy about the existence of the enzyme in this p o u p of anoxygenic phototrophs (Tabita et al., 1974), it was confirmed (Buchanan and Sirevag, 1976; Ormerod and Sirevag, 1983) that ribulose 1,5-bisdiphosphate carboxylase is absent and CO, assimilation occurs by the reductive carboxylic acid cycle. Autotrophic carbon assimilation by anoxygenic phototrophic bacteria is slow compared to that determined in the presence of C, dicarboxylic acids (Ormerod and Gest, 1962; Schmidt and Kamen, 1970; Kamf and Pfennig, 1980). Factors governing the regulation of CO, assimilation in anoxygenic phototrophic bacteria were studied using wild-type (Jouanneau and Tabita, 1986,1987; Valle et al., 1988;Hallenbeck et al., 1990a) and mutant bacteria (Hallenbeck et al., 1990b). The molecular and cellular regulation of autotrophic CO, assimilation has been thoroughly discussed (Tabita,1988;Tabita et al., 1990).The enzyme ribulose 1,5-bisphosphate carboxylase/oxygenase is well characterized (Schloss et al., 1979; Gibson and Tabita, 1985, 1987; Falcone and Tabita, 1991) together with the genetic localization and mapping of autotrophic CO, assimilation genes (Muller et al., 1985; Zhu and Kaplan, 1985; Gibson and Tabita, 1987; Hallenbeck and Kaplan, 1988; Valle et al., 1988; Tabita et al., 1990). Apparently all ribulose 1,5-bisphosphate carboxylases studied from anoxygenic phototrophic bacteria possess associated oxygenase activity (McFadden, 1974; Miziorko and Lorimer, 1983).Dioxygen is a competitive inhibitor of C02, and the enzyme catalyzes the formation of one molecule of 2-phosphoglycolate from ribulose 1,5-bisphospate and 0,. Thermostable enzyme and CO, assimilation have been reported recently (Bakurova and Zhukov, 1986a,b; Heda and Madigan, 1988). Ribulose 1,5-bisphosphate carboxylase was inhibited in cells grown phototropically with malate or succinate as carbon sources (Slater and Morris, 1973; Quayle and Pfennig, 1975; Ferguson et al., 1987). However, synthesis of autotrophic enzymes appeared to be constitutive in Ectothiorhodospira sp. ATCC 31751 (Bognar et al., 1982) when grown on acetate. The presence of thiosulfate also significantly affected the carbon assimilation in the purple nonsulfur bacterium R. palustris (Rolls and Lindstrom, 1967).


261

Apart from autotrophic CO, fixation, photoheterotrophic CO, fixation was also shown in anoxygenic phototrophic bacteria as in other heterotrophs (Dijkhuizen and Harder, 1985). In R. rubrum growing photoheterotrophically on L-malate, one of the major pathways of photometabolism of CO, is through glycolic acid (Anderson and Fuller, 1967a). For assimilation of propionate, butyrate [Ormerod, 1956). and glycerol (Pellerin and Gest, 1983) CO, was found essential; thus, the metabolism of reduced organic acids and of CO, fixation by the Calvin cycle enzymes may be interconnected (Ferguson et a]., 1987). Similarly, in Autmutants of R. capsulatus isolated by Willison et al. (19841, both the photoproduction of hydrogen from organic substrates and autotrophic growth are affected. Large amounts of CO, are evolved from organic acids during photoassimilation, and this CO, forms a reservoir .of potential carbon that would increase cell yields and whose loss could be prevented by addition of excess reducing power [Kikuchi et al., 1961; Rolls and Lindstrom, 1967). Assimilation of one-carbon compounds such as CH,OH, HCOOH, CH, , and CO for growth is exceedingly slow. An inducible enzyme formate hydrogen lyase, which catalyzes the formation of CO, and H, from formate, has been reported among many anoxygenic phototrophic bacteria (Qadri and Hoare, 1968; Schon and Voelskow, 1976; Gorrell and Uffen, 1977, 1978; Voelskow and Schon, 1978). The ability to photoassimilate formate, the presence of formate hydrogen lyase, and the presence of hydrogenase all appear to depend on growth of the organism on a medium containing formate (Qadri and Hoare, 1968). The formate hydrogen lyase of anoxygenic phototrophic bacteria appears to have properties similar to that of heterotrophic bacteria (Qadri and Hoare, 1968). Apart from the autotrophic assimilation, heterotrophic (direct) assimilation of formate via the serine pathway, as found in Pseudomonas sp. AM I (Large and Quayle, 1963), must also be present in anoxygenic phototrophic bacteria because R. sphaeroides O.U. 001 with apparently no capability for autotrophic growth (Sasikala and Ramana, 1990a) was found capable of growth with formate as the sole carbon source (Sasikala, 1990). Photosynthetic bacteria growing under CO have been accidentally discovered in enrichment cultures of CO-oxidizing bacteria (Hirsch, 1968), where Rhodopseudomonas sp. was able to grow with 70% CO as the carbon source along with 20% 0, + 1% CO, + 9% N,. Resting cell suspensions of CO-grown cells of Rhodopseudomonas sp. metabolize about 6.7 pmol of CO/mg protein in 1 hour and produce equimolar amounts of CO, and H, [Table IX) (Uffen, 1976). Carbon monoxide dehydrogenase is assumed to be the enzyme responsible for CO oxidation in anoxygenic phototrophic bacteria (Bonam and Ludden, 1987; Bonam

TABLE M CARFION ASSIMILATION BY ANOXYGENIC PHOTOTROPHIC BACTERIA

Reaction

Reference

I. Autotrophic A. Photoautotrophic 1. COZ 2. COZ

+ 2Hz+ (CHzO) + HzO + 2 HzS+ ((3320)+ HzO + 2 S

Stanier et al. (1959) Stanier et al. (1959), Fuller et al. (1961)

Chemoautotrophic (dark) 17 Na,S20, + 32 CO, + 45 H,O+ 8 (GH,O,) C. Formate assimilation

B.

HCOOH + Hz

+ COz

+

+ 34 NaHSO,

Kampf and Pfennig (1980)

Schon and Voelskow (1976)

(MzO)

D. Methanol assimilation 2CHsOH + COz + 3 (CHzO) + HzO E. Carbon monoxide assimilation CO

+ HzO+

COz

Quayle and Pfennig (1975)

+ Hz

Uffen (19761, Uffen et al. (1990)

F. Mixotrophic 19 Na,S,O, + 32 CO, + 32 CHsCOOH + 39H20+ 24 (GH,O,) II. Organic carbon assimilation A. Dark 1. Endogenous (aerobic) (GH,oOs). + n HzO + 311s- (G&OZ)”+ 2n CO, + 3n H,S 2. Exogenous a. Aerobic G&Os + Oz -+ 2(CHZO) + ZCOZ + HzO b. Anaerobic 18 C3H,03 + 16Hz0-+ 17 CH,COOH

B.

+ 38 NaHSO,

+ 14 H, + 16 C G + 1C,&O, + 1HCOOH

Kampf and Pfennig (1980)

van Gemerdan (1968)

Morita (1961)

Schon and Voelskow (1976)

Light 1. Anaerobic a. Presence of N

G&.os-

c3&03

+ coz

Bregoff and h e n (1952)

b. Absence of N G&Os + 3HzO+4COz + 6H2 2. Aerobic GJ&O, + 0.5 O,-*2.5 (CH,O) 1.5 COz + 0.5 H,O

+

Ormerod and Gest (1962) Morita (1961)


263

et al., 1984,1988,1989),and the enzyme is similar to that of homoacetate-fermenting bacteria (Diekert et al., 1979;Drake et al., 1980;Ragsdale et al., 1983).Carbon monoxide dehydrogenases isolated from anaerobic bacteria are nickel dependent (Diekert et al., 1979;Ragsdale et al., 1983; Bonam and Ludden, 1987), oxygen labile (Bonam et al., 1984),act optimally at pH 8.4 (Ragsdale et al., 1983),and differ from those isolated from aerobic bacteria in having molybdenum and being oxygen stable (Meyer and Schlegel, 1980; Kim and Hegeman, 1981; Meyer, 1982). Recently, Ludden and co-workers (Bonam et al., 1989) observed in R. rubrum ATCC 11170 that CO induces two enzymatic activities, CO dehydrogenase and a CO-insensitive hydrogenase, which appear to function together to accomplish the oxidation of CO to CO, and H,. Both enzymes were activated by 0, in vivo and in vitro, and the synthesis of CO dehydrogenase is shown to be inhibited by oxygen. Apart from R. gelatinosus (Wakim and Uffen, 1983; Champine and Uffen, 1987) and R. rubrum (Bonam et a]., 1989),there are no reports about the assimilation of CO and the related enzymes from other anoxygenic phototrophic bacteria. This area needs further study in terms of both basic (biochemistry) and applied aspects (using them as CO depollutants) as CO is a natural by-product in a variety of biological and chemical reactions that occur in nature (Willison et al., 1970; Uffen, 1976).

Yet another C, carbon compound, methanol, is utilized by a few strains (Quayle and Pfennig, 1975;Douthit and Pfennig, 1976;Sahm et al., 1976;Bamforth and Quayle, 1978;Siefert and Pfennig, 1979),and strains of R. acidophila could tolerate methanol concentrations of 15-250 mM (Quayle and Pfennig, 1975). Growth on methanol was dependent on the simultaneous presence of bicarbonate (Quayle and Pfennig, 1975;Bamforth and Quayle, 1978;Siefert and Pfennig, 1979; Fujii et al., 1982)which functions as an electron acceptor. Rhodopseudomonas acidophila oxidizes methanol to CO, with methanol dehydrogenase and uses the reducing equivalents for CO, fixation via the ribulose bisphosphate cycle (Sahm et al., 1976).The methanol dehydrogenase enzyme resembles numerous previously reported methanol dehydrogenases from methylotrophic organisms (Bamforth and Quayle, 1978).

Unlike autotrophic carbon assimilation where CO, is assimilated either by the pentose phosphate pathway or by the reductive tricarboxylic acid cycle, organic acids are assimilated by the citric acid cycle (Gest et al., 1962;Ormerod and Gest 1962;Beatty and Gest, 1981;Gest, 1972) as confirmed by fluoroacetate inhibition studies. Catabolism of

264

K.SASIKALA ET AL.

organic acids depends on the nutritional status of the organism and the conditions under which it is assimilated. By and large, catabolic assimilation of organic acids in the absence of combined nitrogen or N, results in the formation of H, and CO, (Siegel and Kamen, 1951;Gest et al., 1962;Ormerod and Gest, 1962) via the anaerobic light-dependent citric acid cycle (Gest et al., 1962).However, in the presence of combined nitrogen or N, in light, the citric acid cycle does not operate regularly; instead, coupled oxidation-reduction reactions occur between carboxylic acids of oxidatively different levels (Kikuchi et al., 1961, 1963)resulting in CO, and a C3 compound (Table IX) which is assimilated as carbohydrate (Bregoff and Kamen, 1952;Ormerod and Gest, 1962;Voelskow and Schon, 1978). A few nitrogen compounds like glutamine, peptone, casamino acids, and urea are assimilated as both carbon and nitrogen sources (Bast, 1977,1986).Attempts to produce hydrogen by resting cells suspended in a strictly inorganic medium either failed (Bregoff and Kamen, 1952) or, at best, very low levels of hydrogen evolution occurred (Siegel and Kamen, 1951;Sirevag, 1975;Hillmer and Gest, 1977a;Sasikala et a]., 1990b);endogenous poly-P-hydroxybutyric acid fermentation occurred. The endogenous fermentation of washed cells of Chromatium sp. strain D grown on an inorganic medium containing thiosulfate produced H,S, CO,, and acetate; however, cells grown with H, as the H donor produced CO,, acetate, and H, (Hendley, 1955). Although anoxygenic phototrophic bacteria prefer organic acids for their growth, they can also metabolize carbohydrates, including simple sugars (Szymona and Doudoroff, 1960;Eidels and Preiss, 1970;Conrad and Schlegel, 1974;Nishizawa et al., 1974;Macler et al., 1979;Schultz and Weaver, 1982;Sasaki et al., 1985) and complex sugars like starch (Buranakarl et al., 1985;Singh eta]., 1990).Rhodobacter capsulatus and R. sphaeroides could grow phototrophically and also chemotrophically (aerobically in darkness) with glucose and fructose; R. sphaeroides metabolizes glucose via the Entner-Doudoroff pathway under both growth conditions but metabolizes fructose via the Entner-Doudoroff pathway under chemotrophic conditions and predominantly via the Emden-Meyerhof pathway under phototrophic growth conditions (Conrad and Schlegel, 1977).The metabolism by which carbohydrates are assimilated has been widely discussed (Szymona and Doudoroff, 1960; Eidels and Preiss, 1970;Conrad and Schlegel, 1974, 1977).Extracellular products of R. sphaeroides grown on glucose weregluconic acid (Kitamura, 1972;Nishizawa et al., 1974),a-keto acids, and a polysaccharidelike substance (Nishizawa et al., 19741. Alcohols like methanol, ethanol, n-propanol, and n-butanol have


265

been shown to be assimilated by an NAD-linked alcohol dehydrogenase (Quayle and Pfennig, 1975; Siefert and Pfennig, 1979; Fujii et al., 1982, 1983, 1987; Hirayama et al., 1986). Anoxygenic phototrophic bacteria cannot metabolize alcohols in the absence of suitable electron acceptors such as bicarbonate under light conditions (Fujii et al., 1987);however, Rhodopseudomonas sp. 7 could grow in an alcohol-containing medium without bicarbonate when it was evolving hydrogen and carbon dioxide (Fujii et al., 1983). Reports on hydrogen production from alcohols are limited; though Rhodopseudomonas sp. 7 is known to possess a constitutive NAD-linked alcohol dehydrogenase, it is not enough to explain the hydrogen evolution mechanism from media containing an alcohol (Fujii et al., 1987). Long-chain fatty acids like linoleic, cisvaccenic, oleic, linolenic, palmitoleic acid, myristic, myristoleic, palmitic, and stearic acid were reported to be assimilated by these bacteria (Campbell and Luecking, 1983). Aromatic carbon compounds are assimilated by a few anoxygenic phototrophic bacteria (Proctor and Scher, 1960; Pfennig et al., 1965; Dutton and Evans, 1969, 1978; Pfennig, 1978; Harwood and Gibson, 1986, 1988; Madigan and Gest, 1988; Kamal and Wyndham, 1990; Wright and Madigan, 1991) via an alternate metabolic pathway from the normal aerobidanaerobic route, which results in the formation of pimelate when benzoate is the substrate for R. palustris. Their dissimilation may vary under aerobic and anaerobic conditions (Harwood and Gibson, 1988). The enzymes involved in the photometabolism of aromatic acids are inducible and appear to lack substrate specificity (Dutton and Evans, 1969). It was reported that R. acidophila M402 (Yamanaka et al., 1991) can utilize aromatic alcohols and aldehydes via their respective dehydrogenases. Benzoate is used as a photoheterotrophic growth substrate by R. vannielii (Wright and Madian, 1991). Assimilation of carbon compounds in the dark depends on the substrate used. Dark anaerobic growth with R. rubrum and R. capsulatus was possible either by a strict mixed-acid fermentation of sugars or in the presence of an appropriate electron acceptor via energy-linked anaerobic respiration (Schultz and Weaver, 1982). It was reported that dark anaerobic growth on sugars by R. capsulatus was dependent on accessory oxidants such as dimethyl sulfoxide or trimethylamine Noxide (Yen and Marrs, 1977; Madigan and Gest, 1978), where the required oxidant functions only as an electron sink to permit the management of fermentative redox balance, rather than as a terminal electron acceptor necessary for electron transport-driven phosphorylation (Madigan et al., 1980). However, Schultz and Weaver (1982) observed that

266

K. SASIKALA ET AL.

R. rubrum G9 and R. capsulatus B10 were able to grow fermentatively on sugars in the absence of added oxidants, but required the presence of bicarbonate before fermentative growth could begin. Dark anaerobic fermentation of fructose by R. rubrum was shown to produce succinate, acetate, propionate, formate, hydrogen, and carbon dioxide; however, R. capsulatus produced lactate, acetate, succinate, hydrogen, and carbon dioxide (Schultz and Weaver, 1982). Nonfermentable substrates like succinate, malate, and acetate support dark anaerobic growth only in the presence of an electron acceptor such as dimethyl sulfoxide or trimethylamine oxide (Schultz and Weaver, 1982). Dark anaerobic assimilation of pyruvic acid is catalyzed by pyruvate-formate lyase, the key enzyme helping in the conversion of pyruvate to acetate and formate as major products and to a lesser extent to CO, and propionate (Schon and Biedermann, 1973;Jungermann and Schon, 1974;Schon and Voelskow, 1976).Formate is further metabolized to H, and CO, (Schon and Voelskow, 1976)

-

Pyruvate propionate + acetate Formate -. Hz+ COz

+ formate

Pyruvate-formate lyase activity could not be demonstrated in anaerobic photosynthesizing cells of R. rubrum Ha (Jungermann and Schon, 1974);however, the formate hydrogen lyase of R. rubrum mutant C functioned equally well under anaerobic light or dark conditions (Gorre11 and Uffen, 1978). The stoichiometric formation of various end products of the dark fermentation of pyruvate has been worked out (Schon and Voelskow, 1976), and gas production from formate catalyzed by crude protein extracts of A. rubrum was optimum at pH 6.5-6.7 (Gorrell and Uffen, 1977). Experiments of Willison et al. (1984)on photohydrogen production by Aut- mutants of R. capsulatus indicated that there is an important relationship between photoproduction of hydrogen by nitrogenase and the pathways of carbon assimilation in the cell. It appears that carbon assimilation is an important factor for hydrogen production since preadaptation of carbon-assimilating enzymes was found to have a determining role. This is supported by (1)the inability of resting cells of acetate-grown R. palustris to produce hydrogen from formate (Qadri and Hoare, 1968),(2)the adaptive nature of the enzyme system for hydrogen formation as indicated by kinetic studies of growth and excretion of fermentable products (Schon and Biedermann, 1973), (3) the influence of preculture substrate on the pyruvate-formate lyase activity (Voelskow and Schon, 1978),(4)the inducible nature of the carbon assimilation enzymes (Qadri and Hoare, 1968;Dutton and Evans, 1969;


267

Hillmer and Gest, 1977b), and (5) the metabolic control of exogenous carbon and nitrogen compounds on the reductive pentose phosphate and tricarboxylic acid enzymes (Anderson and Fuller, 1967b). An efficient and effective carbon assimilation system is very important for achieving high hydrogen production values. Although degradation pathways are different for the various substrates, there are common parts like the tricarboxylic acid or glycolate cycle, and a strain with high activity on one substrate would have efficient degradation pathways (Ma0 et al., 1986). VII. Advances in Hydrogen Production Technologies Using Anoxygenic Phototrophic Bacteria

A. UTILIZATIONOF WASTEWATERS The search for abundantly available, inexpensive sources of electron donors for hydrogen photoproduction is the focal area of research for practical exploitation of the process. Organic wastes of various sources with low combined nitrogen contents are ideal substrates for hydrogen photoproduction with the advantage of negative costs corresponding to purification expenses. Wastes used as electron donors for photoproduction of hydrogen include sugar refinery waste (Vincenzini et al., 1981,1982b; Bolliger eta]., 1985),straw paper mill effluent (Vincenzini et al., 1982b), distillery wastewaters (Sasikala et al., 1992; Vatsala and Ramasamy, 1989), lactic acid-containing wastes (Zurrer and Bachofen, 1979; Sasikala et ol., 1991a), and also the clarified slurry of biogas plants (Vrati and Verma, 1983).Agar-entrapped cells of R. palustris and R. molischianum photoproduced hydrogen from sugar refinery waste and straw paper mill effluent for over a month at a rate ranging from 50 to 139 pl HJmg cells dry wthour depending on the organism and on the substrate; in addition, the amount of hydrogen evolved per liter of wastewater ranged from 0.78 to 2.6 liters, with a chemical oxygen demand (COD) conversion to hydrogen of 28-43% (Vincenzini et al., 1982b). Bolliger et al. (1985),while studying hydrogen evolution with sugar refinery wastes serving as the electron donor, observed that naturally adapted organisms enriched and selected from a similar environment proved better for hydrogen production from wastes than did the laboratory strains. Apart from the above, where anoxygenic phototrophic bacteria alone were used for hydrogen photoproduction from wastes, two-stage processes involving heterotrophs in the first stage have also been reported (Vrati and Verma, 1983; Karube et al., 1984). A two-stage fermentation

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of cow dung was proposed by Vrati and Verma (1983).During the first stage, the cow dung is anaerobically fermented; the effluent slurry is separated into a solid residue, which could be used as a feed supplement, and a supernatant liquid fraction, which was used for single cell protein (SCP) and hydrogen photoproduction by R. capsulatus. Alginate-immobilized R. rubrum was used to improve the efficiency of hydrogen production by the heterotrophic bacterium Citrobacter freundii from diluted molasses (Karube and Suzuki, 1988;Karube et al., 1984), where R. rubrum was found to evolve hydrogen (19-31 ml/minute) continuously for over 90 hours from the wastewater of the above fermentation containing organic acids. The hydrogen was supplied to a phosphoric acid fuel cell, producing a stable current from 0.5 to 0.6 A for 6 hours and a power of 0.16-0.18 W. The only commercially exploited practical system of hydrogen photoevolution from wastewaters is that of Kobayashi and Kondo (19841,where from an organic waste solution with a biological oxygen demand (BOD) value higher than 10,000ppm, the bacteria produced more than 150 liters/day/m3 of hydrogen gas. Efforts continue to utilize agricultural wastes, and studies of hydrogen photoproduction from complex polysaccharides like starch (Buranakarl et al., 1985;Singh et al., 1990)and cellulose (Vatsala, 1989)have been initiated recently. B. COCULTURES

Work has only recently been initiated on the production of hydrogen by associations of various organisms. Hydrogen evolution by various photosynthetic bacteria associated with the heterotrophs Klebsiella pneumoniae (Weetall et al., 1981),Cellulomonas sp. ATCC 21399 (Odom and Wall, 1983), Clostridium butyricum (Miyake et a]., 1984, 1985), and Pseudomonas flouresence (Sasikala, 1990) and the phototrophs Chlamydomonas reinhardtii (Miyamoto et a]., 1987)and Synechococcus cedrorum (Sasikala, 1990) has been reported. Immobilized cells of an apparently contaminated (with Klebsiella pneumoniae) culture of R. rubrum (Weetall et al., 1981) were found to photoproduce hydrogen from glucose (hydrolyzed cellulose) and raw sawdust in single-pass and recycle reactors. The mixed culture resulted in better use of glucose, and more than 100% efficiencies were observed with the cellulose hydrolyzate (calculated as percentage efficiency assuming 6.0 mol hydrogen per mole of glucose), apparently because of utilization of other substrates in the hydrolyzate. The combination of photosynthetic bacteria with anaerobic bacteria


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could provide a system for hydrogen photoproduction from carbohydrates which are either poorly utilized (glucose) or not utilized (cellulose) by phototrophic bacteria. In such a system, anaerobic bacteria produce intermediates as organic acids, which are then converted to hydrogen by photosynthetic bacteria. For coupling cellulose utilization to photoproduction of hydrogen, cocultures of Cellulomonas sp. ATCC 21399 and R. capsulatus BlOO were employed (Odom and Wall, 1983). Cellulomonas produced formate, acetate, lactate, and small amounts of succinate, apart from the considerable amounts of reducing sugars, presumably cellobiose, which were used by R. capsulatus for photoproduction of hydrogen. Miyake et al. (1984,1985) have studied photoproduction of hydrogen from glucose by a coculture of Clostridium butyricum and Rhodopseudomonas sp. RV. The C. butyricum produced acetate, lactate, and butyrate along with CO, and H,, and butyrate was the major product. This two-step production of hydrogen via butyrate from glucose showed a high efficiency of 7.0 mol hydrogen from 1 mol of glucose, of which 16% was estimated to be produced by C. butyricum. Concerning energetic aspects (Miyake eta]., 1985), it has been shown that the combined process requires less energy than the reaction carried out by nitrogenase alone (i.e., by Rhodopseudomonas sp. RV), owing to the balanced use of nitrogenase (energy-dependent) and hydrogenase (which is energy independent) from an energy viewpoint. In another system, combining a green alga Chlamydomonas reinhardtii with R. rubrum (Miyamoto et al., 1987),however, only the dark hydrogen evolution capacity of the photosynthetic bacteria was exploited. The process was carried out under light (aerobic) and dark (anaerobic) cycles. In the light phase, the green alga accumulated carbohydrate, which was fermented in the dark period to produce several organic compounds (formate, acetate, ethanol, and glycerol, apart from CO, and H,) among which formate was utilized for dark hydrogen production by the photosynthetic bacterium. Here, the dark fermentative production of hydrogen by the green alga is augmented by the photosynthetic bacterial hydrogen evolution from formate, and elevations of about 4-fold in hydrogen evolution rate and 5-fold in hydrogen molar yield (moles hydrogen per mole glucose) were obtained with the mixed culture compared with that of C. reinhardtii alone. Hydrogen photoproduction by mixed cultures of R. sphaeroides O.U. 001 with the cyanobacterium Synechococcus cedrorum and the heterotrophic bacterium Pseudomonas fluorescens (Sasikala, 1990) showed enhanced hydrogen photoproduction by the mixed cultures. However, such enhancement was found only when the various organisms were immobi-

TABLE X

HYDROGEN PHOTOPRODUCTION INPEIOTOBIOREWTORS

Reference Mignot eta]. ( 1 9 8 9 ~ ) Segers and Vestmete

Reactor volume0 (mu NM 400

Organism R. rubrum 7061 A. rubrum ATCC 11170

Free or immobilized Agar immobilized Free cells

Duration (days) >4.15 40

Total hydrogen produced (LI >0.0175 >3

(1983)

Kim et d.(1981) Stevens et d.(1986) Hirayma et al. (1986)

Karube et d.(1984) Stevens et aJ. (1983) Delachapelle et al. (1991) Vincenzini et d.(1982a) Kim eta]. (1982a) Kim et d.(1987a) von Felten et 01. (1965) Bennett and Weetall (1976) Planchard et al. (1989) Zurrer et al. (1981) Mitsui et aJ. (1985)

750 2500 400 140 40,000 450 11,Ooo 680 33,000 6000 NM

NM 200 1000

NM

A. sphaeroides TN 3 R. sulfidophilus LMG 5202 R. rubrum G 9 BM

R. rubrum IF0 3986 A. rubrum IF0 3986 R. capsulatus R. capsulatus A. molischianum R. sphaeroides B5 A. sphaeroides B6 R. rubrum R. rubrumR. rubrum 7061 A. rubrum Rhodopseudomonas sp. Miami

Free cells Free cells Agar-cellulose fiber immobilized Alginate immobilized Alginate immobilized Free cells Free cells Agarized panel Free cells . Free cells Agar immobilized Agar immobilized Agar immobilized Free cells Agar immobilized

Several days

Alginate immobilized

12

12 3 60 6 3.75 5 10 35 25 46.3 125 150 8.45 >8

1.67 2.975 0.577 0.624 >80 3.667 170 2.8 177.2 125.4 0.8 0.198 0.037 >4.0 0.086 per day

2271

Sasikala et 01. (1992)

NM,Not mentioned.

3600

R. sphaeroides O.U. 001

3.0


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lized individually and used for hydrogen production in mixed cultures in contrast to mixing the cell suspensions first and then immobilizing (coimmobilization), where hydrogen production was inhibited. The cause for such a property has yet to be established since coimmobilized mixed cultures of K.pneumoniae and A. rubrum (Weetall et al., 1981) and C. butyricum and Rhodopseudomonas sp. RV (Miyake et al., 1984) could produce hydrogen. Mixed cultures of anoxygenic phototrophic bacteria with other organisms have been used for other purposes, including biomass production (Odom and Wall, 1983; Noparatnaraporn et al., 1987a),nitrogen fixation (Kobayashi et al., 1981), and augmentation of methane in a biogas process (Vatsala and Balaji, 1987).

C. PHOTOBIOREACTORS An important aspect to be studied in biological hydrogen production for the practical application of anoxygenic phototrophic bacteria as compared to heterotrophs is the reactor design, which is complicated by the requirement for light. However, this aspect has been only sparingly studied, with only a few reports available at the laboratory scale, ranging from a few hundred to a few thousand milliliters (Table X). The two important considerations for reactor design are providing adequate homogeneous illumination and reducing the effect of heat generated owing to the illumination. All the reactors designed so far have the source of illumination outside the reaction vessel, and hence the material out of which it is made is transparent glass (Mignot et al., 1989c; Vincenzini et al., 1981; Sasikala et al., 1992), acrylate (Stevens et al., 1983; Kim et al., 1982a; Karube et al., 1984), or polycarbonate (Zurrer et al., 1981). The geometry of the reactor fundamentally affects the illumination reaching the cells and hence hydrogen photoevolution. The reactors used are mostly rectangular or columnar. Stevens et al. (1983)used an all-glass reactor consisting of a medium chamber between two concentric cylindrical walls with the illumination being provided by a bulb at the center (Fig. 1). Such an arrangement provides better illumination as compared to illumination from one or a few sides (von Felten et al., 1985; Stevens et al., 1986; Planchard et al., 1989; Sasikala et al., 1992); however, the problem of heat generation is more severe. To overcome this inhibiting thermic effect, an innovative approach of using optical fibers to guide light to the reactor was used successfully by Mignot et al. (1989a,b,c) in a patented design (Junter et al., 1989) of a bioreactor based on a tubular gel layer/microporous membrane structure around an inner optical fiber illumination device, which ensures the biocatalytic layer both ho-

‘1V.L3V I m I s V s ‘>I

ZLZ


U

u

u

18

17

273

U 16

FIG. 2. New type of immobilized-cell photobioreactor with internal illumination by optical fibers. [After Mignot et al. (1989c).] (1)Optic-fiber barrel, (2) inner glass cylinder, (3) immobilized-cell agar layer, (4) outer cylindrical stainless steel grid, (5) microporous membrane, (6) assembling pin, (7) tank, ( 8 ) reactor wall, (9) nut, (10) reactor cap, (11) stainless steel plate, (12)bottom plate, (13)bracket, (14) pH electrode, (15) temperature probe, (16) heating resistance, (17)nutrient medium inlet, (18) nutrient medium outlet, (19) rubber band, (20) safety valve, ( 2 3 ) gas outlet, (22) argon inlet.

plete reactor into a thermostatted water bath (Fig. 6).Zurrer et al. (1981) used a cooling coil that limits the temperature to 30°C in a vessel above the transparent panel of the reactor, and an energy saving circulation of the cell suspension is achieved. Economically, however, energy cannot be expended to heat or cool the reactor because the process is not very efficient, so the reactor must be designed to maintain a suitable temperature (Herlevich and Karpuk, 1982).The pH of the reaction is controlled by on-line monitoring and adjusting by an automatic pH controller (Hirayama et al., 1986,see Fig. 4; Stevens et a]., 1986) or by buffering the medium (Mignot et a]., 1989c;Stevens et al., 1986).Stirring is another important factor because it was proved that it enhances not only total hydrogen evolution but also the conversion efficiency of substrate to hydrogen (Kim et al., 1981).A magnetic stirrer has been

K.SASIKALA ET AL.

2 74 3

A

0

0

FIG.3. Biosolar reactor used for outdoor hydrogen production by R. sphaeroides. The reactor was made of polyacrylate of 8 mm thickness. It contained 33 liters of culture medium and was equipped with a gas outlet, a manometer, a pH electrode, and a medium temperature senso: [After Kim et al. (1982a).] (A) Cross section, (B) longitudinal aspect. (1)pH electrode, (2) temperature sensor, (3) gas outlet, (4) manometer for measuring internal pressure, (5)bovine venous injection needle for sampling, (6) reinforcement plate.

used for this process (Fig. 7) (Kim et al., 1981);however, because the process involves gas evolution, agitation is created automatically, which is enhanced by scaling up and in larger vessels (33 liters) was found to cause enough agitation to increase yields (Kim et al., 1982a). Both free (Zurrer and Bachofen, 1982; Kim et al., 1982a,b, 1987a,b; Stevens et al., 1986) and immobilized cells (Sasikala et al., 1992; von Felton et al., 1985; Planchard et al., 1989; Mitsui et al., 1985; Hirayama et al., 1986; Mignot et al., 1989c) are employed in reactors, and when immobilized cells are used in the form of a panel (Fig. 7) there is the necessity of providing support, which has been achieved by immobilizing on a nylon net (Planchard et al., 1989), on an agarose-coated polyster film (Mitsui et al., 1985), on a plastic slab composed of a series of thin plastic layers (Bennett and Weetall, 1976), or in a loom having little holes along the inner edge (Vincenzini et a]., 1981). Anaerobic conditions are created in the reactor by flushing an inert gas (Ar or He), by leaving very little (Sasikala et al., 1992) or no headspace (Vincenzini et al., 1981),or by using a liquid paraffin layer (Kim et al., 1981, 198713).


275

FIG.4. Apparatus used for continuous hydrogen production by immobilized R. rubrum. The reactor was made of polyacrylate sheets of 2 mm thickness. The vessel (inner size 1.5 x 15 x 18 cm) contained the immobilized cells with the substrate solution and was equipped with a water inlet, a water outlet, a gas outlet, a pH electrode, and a thermometer. [After Hirayama et aJ. (1986).] (1) Reactor vessel, (2) thermometer, (3) water inlet, (4) water outlet, (5) gas outlet, (6) pH electrode, (7)electric light bulb, (8) glass syringe, (9) pH controller, (10)microtubing pump, (11)substrate solution tank, (12) graduated cylinder for storage of treated water.

While comparing a number of reactor designs (tubular, insulated channel, uninsulated channel, and deep pond reactors) in an energy (theoretical) analysis to examine system requirements for the application of an anoxygenic phototrophic bacterial process to large-scale hydrogen production, Herlevich et aJ. (1982) found that a deep tank reactor was the most attractive. Their studies revealed that the production of hydrogen from anoxygenic phototrophic bacteria seemed technically and economically feasible from an engineering viewpoint. The conceptual design of the deep tank reactor with a nominal production capacity of 28,000 m3/day (1 x los ft3/day)showed promise, warranting further engineering research which is being continued with a computer program called SOLBUG (Herlevich, 1983);however, such a system has not been used practically. Outdoor hydrogen photoevolution was demonstrated in biosolar reactors (Fig. 3) for 25 days from lactate (Kim et al., 1982a) and for several days from orange-processing wastewater (Mitsui et al., 1985). Kobayashi and Ye (1986) developed a technique for outdoor production of

FIG. 5. All-glass column reactor used for hydrogen production from distillery waste by immobilized R. sphaeroides. The reactor (3.6 liters, 72 x 8 cm) was provided with thermometer and pH electrode at the middle, and at the bottom there was a provision to withdraw the reaction mixture. Hydrogen was collected by reversible displacement after passing through three trappers containing 30% KOH. [After Sasikala et al. (1992).]

FIG.6. Experimental setup used by von Felten et al. (1985) for hydrogen evolution by immobilized R. rubrum. (1) Reaction vessel, in a thermostatted water bath, illuminated by tungsten lamps, (2) substrate supply, (3) time controlled pump for medium supply to the reactor, (4) medium overflow, (5) gas sampling vessel for volume determination, (6) overpressure with argon for anaerobic conditions,

277


2 1

3

FIG.7. Reactor used for hydrogen production by planar agar matrix bounded by a microporus membrane filter containing immobilized R. rubrum. [After Planchard et al. (1989).](1)Reactor vessel, (2) gas outlet, (3) measuring test tube for gas volume determination, (4) air flow control (blower), (5) thermostatted water flow, (6) 150-W tungsten incandescent bulb, (7) magnetic stirring, (8) light-transparent plate, (9) bolt, (10) microporous membrane filter, (11)agar layer entrapping viable A. rubrum cells.

278

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hydrogen on a large scale, producing only 150 liters hydrogen gadday/ m3 using a pure culture of anoxygenic phototrophic bacteria. The output was tripled (500 liters/day/m3) on culturing with Bacillus megaterium, and 1000 liters/day/m3 was produced when anoxygenic phototrophic bacteria were cocultured immediately after sufficient growth of aerobic heterotrophs collected from a natural source. During long-term outdoor hydrogen photoproduction, the problem of algal growth in the reactors can be controlled by the use of algal inhibitors like “chloroxuron,” which had no detrimental effect on anoxygenic phototrophic bacterial growth and nitrogenase activity (Segers and Verstraete, 1985). Even in large-scale operations in bioreactors, the hydrogen was not stored or used for any purpose, except in the work of Karube et al. (1984) where hydrogen produced by immobilized R. rubrum was passed through a phosphoric acid fuel cell (Fig. 8) and a stable current

02

FIG.8. Large-scale bacterial fuel cell using immobilized R.rubrum. Two large acrylate bioreactors (20 liters) with about 18 kg of immobilized A. rubrum were operated at pH 6.87.0, 30°C, and a light intensity of 5000 lux with wastewater continuously transferred to the reactors at a flow rate of 2.0 litershour. Hydrogen produced by the reactors was transferred to the anode of a phosphoric acid fuel cell system at 19-31 ml/min. Oxygen was also transferred to a cathode at 0.1 litedminute. [After Karube et al. (1984).] (1,2) Bioreactors for immobilized R. rubrum, (3-5) hydrogen reservoirs, (6) KOH solution, (7) water, (8) phosphoric acid fuel cell, (9) variable electronic load, (10) heater with flow indicator (FI) and temperature control (TC).


279

of 0.5-0.6 A was obtained for 6 hours. Similar reports of current generation were observed for the cyanobacterium Anabaena N 7363 (Karube et al., 1986) and the heterotrophic bacterium Clostridium butyricum. The stoichiometry of hydrogen to carbon dioxide produced is an important factor to be considered for biofuel cell applications because carbon dioxide is known to impede function (Appley and Fonlkes, 1989). Little work has been carried out on the ratios of hydrogen to carbon dioxide production, and, i n practice, theoretical stoichiometric ratios of hydrogen and carbon dioxide are not observed, with the concentration of hydrogen in the evolved gas mixture reaching 80 to almost 100% (Segers and Verstraete, 1983; Margaritis and Vogrinetz, 1983), depending on the illumination, stirring (Kim et al., 1982a), duration of incubation (Kim et al., 1987b), pH, and pressure (Zurrer and Bachofen, 1982). Phototrophs can also act as anodic photoelectrodes on continuous illumination, as observed with the cyanobacterium Mastigocladus laminosus, for periods of time adequate for use in a konventional photoelectrochemical cell producing about 80- 100 nA current (Ochiai et al., 1980); however, such an attempt has not yet been reported for anoxygenic phototrophic bacteria.

VIII. Other Uses of Anoxygenic Phototrophic Bacteria Apart from photoproduction of hydrogen, other potentially exploitable attributes of anoxygenic phototrophic bacteria include single cell protein (SCP) production and wastewater treatment which may become useful by-products of the process and enhance the overall feasibility and economical viability for practical applications. The biomass of photosynthetic bacteria is very rich in protein content (60-70% of dry wt; Vrati and Verma, 1983; Vrati, 1984; Sasaki et al., 1981; Shipman et al., 1975; Noparatnaraporn et al., 1987a,b) with essential amino acids comparable to those of soy bean and egg proteins (Sasaki et al., 1981; Shipman et al., 1975) or higher than that of other SCP (Noparatnaraporn et al., 1987b) and meat (Driessens et al., 1987). The high vitamin content (Noparatnaraporn et al., 1987a; Kobayashi and Kurata, 1978) also makes it an ideal feed supplement for livestock and fish. It was indeed shown that both the quality and quantity of egg laying by hens was enhanced (Kobayashi, 1976; Kobayashi and Kurata, 1978; Hirayama and Ishigaki, 1988) and gill corruption disease of lobster and Marek’s disease of egg-laying hens were suppressed (Kobayashi and Konda, 1984) by feed supplementation with anoxygenic phototrophic bacteria. Application in pisciculture (Kobayashi, 1976; Noparatnaraporn et al., 1987b) and horticulture (Kobayashi and Kurata,11978)were also found to be beneficial.

280

K. SASIKALA ET AL.

Various agricultural and industrial wastes were used as substrates for SCP production (Suhaimi et al., 1988; Shipman et al., 1975; Kobayashi and Kurata, 1978; Vrati and Verma, 1983; Vrati, 1984; Sawada et al., 1977; Noparatnaraporn et al., 1983; Sasaki et al., 1981; Suhaimi et al., 1988; Hiraishi et al., 1989). Though conditions for SCP production have been optimized (Nopartnaraporn et al., 1983; Driessens et al., 1987; Sawada and Rogers, 1977) and cell mass can be used as a multipurpose animal feed supplement, for practical applications the cost must be decreased, and long-term toxicity tests need to be performed (Noparatnaraporn et al., 1987b). Other products of interest from anoxygenic phototrophic bacteria include vitamin B,, (Vrati and Verma, 1983; Sasaki and Nagai, 1979; Noparatnaraporn et al., 1986; Hirayama and Katsuta, 1988),ubiquinone Qlo (ubielecarenone) (Kobayashi and Kondo, 1984; Carr and Excell, 1965; Sasaki and Nagai, 1979), bacteriocins (Guest, 19741, 5-aminolevulinic acid (ALA) (Sasaki et al., 1987, 1989), some antiviral substances (Kobayashi et al., 1978; Kobayashi and Kondo, 1984; Hirotani et al., 1991), hopanoid having toxicity against mouse leukemia cells (Nagumo et al., 1991), antibiotic (Burgess et al., 1991), ammonia leaching (Sasikala and Ramana, 1990b),augmentation of methane content in biogas (Vatsala and Balaji, 1987), and metal ion extraction (Vatsala, 1987b). Anoxygenic phototrophic bacteria are known to degrade toxic compounds (dimethylnitrosamine), and their role in waste degradation in nature is well established (Kobayashi et al., 1977). They are being further exploited for BOD/COD reduction in wastes of various origins (Sawada and Rogers, 1977; Kobayashi et a]., 1978; Zurrer and Bachofen, 1979; Mitsui et al., 1985; Hiraishi et al., 1989; Vatsala and Ramasamy, 1989). The advantages of using anoxygenic phototrophic bacteria instead of an activated sludge process for waste treatment are that, unlike the activated sludge process where an additional sludge disposal problem is created, photosynthetic bacterial treatment of wastes produces a bacterial biomass rich in proteins and many other valuable organic substances (Balloni et al., 1980);moreover, because dilution of wastewater is not required, the process can be used in areas of water scarcity and the area required for the treatment facilities is decreased (Kobayashi and Yoshida, 1983). Treatment of wastes by anoxygenic phototrophic bacteria is being carried out along the following lines, which may also be coupled with biomass and/or hydrogen photoproduction: (1)onestage process where a given species of anoxygenic phototrophic bacteria is selectively used on a given waste (Vincenzini et al., 1982b; Vat-


281

sala and Ramasamy, 1989); (2)two-stage process with two subsequent treatment steps where (a) the anoxygenic phototrophic bacterial stage either precedes the cyanobacterial (Balloni et al., 1980)or follows the heterotrophic bacterial stage (Sawada et al., 1977) or (b) a photosynthetic bacterial stage is followed by an activated sludge process after dilution (Sasaki et a]., 1981);(3)the involvement of sequential growth of microorganisms in a multistage process (Kobayashi and Yoshida, 1983).Commercialization of processes for purification of wastes of various origins by phototrophic bacteria began about 20 years ago, and purification plants are being operated in many countries (Kobayashi and Kondo, 1984). IX. Conclusion

Biological hydrogen evolution is still in an exploratory stage in bioenergy production research, unlike methane (biogas) and alcohol production. There is enough justification to carry out further work which should now concentrate on process development rather than on strain selection. For this futuristic research, a multi- and interdisciplinary approach is the need of the day. ACKNOWLEDGMENTS The authors are grateful to Drs. P. M. Vignais and A. Colbeau (Laboratoire de Biochimie Microbienne, CNRG, Grenoble, France) for critical reading of the manuscript and for helpful advice and to Dr. Johannes F. Imhoff (Institut fur Mikrobiologie and Biotechnologie, Rheinische Friedrich-Wilhelms-Universitat Bonn, Bonn, Germany) for helpful advice. K. s.and Ch. V. R. thank the CSIR, New Delhi, for the award of Research Associateships. P.R.R. acknowledges the financial support of ICAR, New Delhi.

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Albracht, S . P. J., Kalkman, M. L., and Slater, E. C. (1983).Biochim. Biophys. Acta 724, 309-316.

Albracht, S. P.J., van der Zwaan, J. D., and Fontijn, R. D. (1984).Biochim. Biophys. Acta 766,245-258.

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Albracht, S. P. J., van der Zwaan, J. W., Fontijn, R. D., and Slater, E. C. (1986).In “Frontiers in Bioinorganic Chemistry” (A. V. Xavier, ed.), pp. 11-19. Verlagsgesellschaft, Weinhein, Germany.

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INDEX

Air, solid-state fermentation and,

A

107-108,110,113-114

Acetate, anoxygenic phototrophic bacteria and, carbon assimilation, 264,266 hydrogen production, 224-227,236,269 Acetone, anoxygenic phototrophic bacteria and, 251 Acetylcholine, antibody technologies and,

Alcohol, anoxygenic phototrophic bacteria and, 221,224,227,264-265 Aldehydes, genetically engineered microorganisms and, 9 Algae, anoxygenic phototrQphicbacteria and, 212,259,269,278 Alkaline phosphatase antibody technologies and, 185,189-191 genetically engineered microorganisms and, 3,94 methods of study, 16,36-38 results, 69,78 Aluminum hydroxide, antibody technologies and, 153-154 Amino acids anoxygenic phototrophic bacteria and,

156

Acetylene, genetically engineered microorganisms and, 94 N-Acetylglucosamine, solid-state fermentation and, 130 Acid phosphatases, genetically engineered microorganisms and, 3,94 methods of study, 16, 36-38 results, 69,78 Acquired immunodeficiency syndrome (AIDS),antibody technologies and,

279

classification, 216,219 enzymes, 243,255 hydrogen production, 221, 230 antibody technologies and, 151-152,

174-175

Actinomycetes, genetically engineered microorganisms and, 56 Adjuvants, antibody technologies and, 153-155.182

ADP, anoxygenic phototrophic bacteria and, 246 ADP-ribosylation anoxygenic phototrophic bacteria and, 245

antibody technologies and, 184 Aeration, solid-state fermentation and,

165,177,189

enzyme immunoassay, 189 immunotoxins, 184 radiolabels, 186 recombinant antibodies, 177 recombinant DNA, 192-194 Amino acids medium, genetically engineered microorganisms and, 29-30 Aminopterin, antibody technologies and,

107-108,111,114

Affinity chromatography, antibody technologies and, 160-161 Affinity purification, antibody technologies and, 152,192 AIDS (Acquired immunodeficiency syndrome], antibody technologies and,

161

Ammonia, anoxygenic phototrophic bacteria and, 230,241,246,249 Ammonification, genetically engineered microorganisms and, 79 Ammonium anoxygenic phototrophic bacteria and,

174-1 75

223,229,242,246-247,253 297

298

INDEX

genetically engineered microorganisms and, 62,87 Ammonium chloride, anoxygenic phototrophic bacteria and, 223,229-230 Ammonium-oxidizer broth, genetically engineered microorganisms and, 25 Amylase, solid-statefermentation and, 103

Anabaena, hydrogen production technology and, 279 Anabaena variabilis, hydrogen productian technology and, 248 Anaerobiosis anoxygenic phototrophic bacteria and carbon assimilation, 259, 262-266 classification, 216 enzymes, 243,255 hydrogen metabolism, 219-220 hydrogen production, 220, 225,229, 266-269,276

solid-statefermentation and, 103, 125 Anion-exchangechromatography, antibody technologies and, 159 Anoxygenic phototrophic bacteria, 211-213,281

carbon assimilation, 259-267 classiRcation,213-217, 219 enzymes hydrogenase, 249-259 nitrogenase, 240-249 hydrogen metabolism, 216-220 hydrogen production, 220-221 electron donors, 221-224 immobilization technology, 235-240 optimization of process, 229-235 substrate conversion efficiency, 224-229

hydrogen production technology cocultures, 266-269.271 photobioreactors, 270-279 wastewaters, 267-268 uses, 279-281

Anti-idiotype antibody technologies, 155-157.182

Antibiotic resistance, genetically engineered microorganisms and, 36 Antibiotic-resistant phenotypes, genetically engineered microorganisms and, 32-35

Antibiotics anoxygenic phototrophic bacteria and, 260

antibody technologies and, 195 genetically engineered microorganisms and, 69,91 Antibodies, anoxygenic phototrophic bacteria and, 251 Antibody-dependent cellular cytotoxicity, antibody technologies and, 171,175, 181

Antibody technologies affinity chromatography, 160-161 enzyme immunoassay, 167-191 future, 194-195 high-performance liquid chromatography, 158-159 history, 150- 151 immunofluorescence, 186- 187 immunoglobulin-binding proteins, 157-156

immunotherapy, 180 cancer, 181-182 immunoglobulin fragments, 160-181 transplantation, 161 immunotoxins, 182-163, 185 Pseudomonas exotoxin, 184-185 ricin, 163-184 monoclonal antibodies, 161-162 bispecificity, 166-167 catalytic antibodies, 167-170 cell fusion, 162 hybridomas, 163-164 nonmurine, 164-165 specificity, 165-166 polyclonal antibodies adjuvants, 153-155 delivery, 155 idiotypes, 155-157 lipid-derivatized antibodies, 155 synthetic peptides, 151-153 precipitation, 158 radiolabels, 186 recombinant antibodies eukaryotes, 171-172 heterochimeras, 174- 175 homochimeras, 175-177 libraries, 172-174 mutagenesis, 174

299

INDEX prokaryotes, 170-1 71 single chain, 177-179 size, 179-180 recombinant DNA affinity purification, 192 epitope mapping, 192-194 immunoscreening, 191-192 Antigens, antibody technologies and, 150, 187,192,194

enzyme immunoassay, 187-189,191 immunotherapy, 180- 182 monoclonal antibodies, 162,164-167, 169- 170

polyclonal antibodies, 151-156 recent developments, 160 recombinant antibodies, 174-177, 179- 180

Antimicrobial agents, genetically engineered microorganisms and, 4,92 Arylsulfatases, genetically engineered microorganisms and, 3,94 methods of study, 16,36,38-40 results, 69 Aspergillus niger, solid-state fermentation and, 120 Aspergillus ochraceus, solid-state fermentation and, 113 Aspergillus oryzae, solid-state fermentation and, 119,132 ATP anoxygenic phototrophic bacteria and, 241-243,245-246,253

genetically engineered microorganisms and, 17 Autoimmune diseases, antibody technologies and, 156,195 Autotrophy, anoxygenic phototrophic bacteria and, 216,252,259-263 Avicel, solid-state fermentation and, 116 Avidin, antibody technologies and, 190 Azotobacter vinelandii, hydrogen production technology and, 247-248

B B cells, antibody technologies and immunotherapy, 182 monoclonal antibodies, 162,164-166

polyclonal antibodies, 152-153, 155 Bacillus megaterium, hydrogen production technology and, 278 Bacillus subtilis, solid-state fermentation and, 132 Bacteria anoxygenic phototrophic, see Anoxygenic phototrophic bacteria antibody technologies and, 194 immunoglobulin-binding proteins, 157-158

immunotherapy, 180 polyclonal antibodies, 154 recent developments, 160 recombinant antibodies, 171,175 recombinant DNA, 193-194 genetically engineered microorganisms and, 4,91,93 methods of study, 11,48 microbial assays, 19-21, 35 nitrogen transformations, 87 results, 56,62,78 soil preparation, 17-18 solid-state fermentation and, 100,113 Barium chloride, genetically engineered microorganisms and, 9-10, 15 Basal medium, genetically engineered microorganisms and, 28-29 Batch reactor, solid-state fermentation and, 112 Beauveria bassiana, solid-state fermentation and, 130,133 Bicarbonate, anoxygenic phototrophic bacteria and, 247,263,265-266 Bin fermentors, solid-state fermentation and, 111 Biochemical phenotypes, genetically engineered microorganisms and, 35.41 Biological oxygen demand, anoxygenic phototrophic bacteria and, 268,280 Bioreactors, see also Photobioreactors antibody technologies and, 164 solid-state fermentation and, 102 experimental measurements, 134,142 heat dissipation, 111-112 mass transfer, 108-111 mathematical modeling, 118,123 physical parameters, 118 Biotin, antibody technologies and, 162,190

INDEX

300 Bispecific antibody technologies, 166-167

Bone marrow, antibody technologies and, 181-182

enzymes, 247 hydrogen metabolism, 219 hydrogen production, 220,223,235, 269,271

Brodyrhizobium japonicum, genetically engineered microorganisms and, 93 Bray’s nitrate-nitrite powder, genetically engineered microorganisms and, 23-24

Butyrate, anoxygenic phototrophic bacteria and, 224-227,269

genetically engineered microorganisms and, 3,6,90,93-94 metabolic activity, 8-16, 52-53 methods of study, 41,49 results, 74 soil enzymes, 62 solid-state fermentation and, 103, 109, 112,130

experimental measurements, C

132-133,140

mathematical modeling, 118-121, Calcitonin, antibody technologies and, 152

Calcium, antibody technologies and, 192 Cancer, antibody technologies and, 182-183,185-186

Cancer immunotherapy, antibody technologies and, 181 Capillary forces, solid-state fermentation and, 113 Carageenan, anoxygenic phototrophic bacteria and, 236, 238-239 Carbodiimides, antibody technologies and, 151-152 Carbohydrate antibody technologies and, 160,186, 189

genetically engineered microorganisms and, 35-36 hydrogen production technology and, 221,227,264,269

solid-state fermentation and, 115,130 Carbon anoxygenic phototrophic bacteria and enzymes, 259-267 hydrogen production, 228, 244, 246-247

genetically engineered microorganisms and, 3 , 1 6 , 4 4 , 5 6 , 9 4 solid-state fermentation and, 100,105 Carbon dioxide anoxygenic phototrophic bacteria and carbon assimilation, 259-261, 264, 266

classification, 216

123

Carbon monoxide, anoxygenic phototrophic bacteria and, 241-242, 257, 261-263

Carbonate, anoxygenic phototrophic bacteria and, 223, 247 Carboxylic acid, anoxygenic phototrophic bacteria and, 260 Carcinoembryonic antigen, antibody technologies and, 176, 184 Cascade method, antibody technologies and, 165 Catalytic antibody technologies, 167-170 CD4, antibody technologies and, 174, 177, 180

cDNA, antibody technologies and, 171, 191-192

Cell fusion, antibody technologies and, 162,166

Cell-mediated immunity, antibody technologies and, 154 Cellulases, genetically engineered microorganisms and, 37 Cellulolytic bacteria, genetically engineered microorganisms and, 56,62 Cellulose anoxygenic phototrophic bacteria and, 223-235,239-240,268

genetically engineered microorganisms and, 21-22,56 solid-state fermentation and, 114-116, 125

Chemotherapy, antibody technologies and, 181-182

301

INDEX

Chimeric antibody technologies, 182 Chitin genetically engineered microorganisms and, 22-23 solid-state fermentation and, 130 Chitinoclastic bacteria, genetically engineered microorganisms and, 56, 62, 78

Chlamydomonas reinhardtii, hydrogen production technology and, 268-269 Chlorobium limicola, hydrogen production technology and, 251 Chlorochromatium aggregatum, hydrogen production technology and, 216 Chlorochromatium glebulum, hydrogen production technology and, 216 Chloroflexus aurantiacus, hydrogen production technology and, 217, 240, 242,244

2-Chloromaleylacetate, genetically engineered microorganisms and, 74 Chorismate mutase, antibody technologies and, 170 Chromatiaceae, hydrogen production technology and, 216-217,220 Chromatium, hydrogen production technology and, 230,232-233,235,264 Chromatium minutissimum, hydrogen production technology and, 220 Chromatium tepidum, hydrogen production technology and, 217 Chromatium vinosum, hydrogen production technology and, 219-220, 250-254,256

Chromatofocusing, antibody technologies and, 159 Chromatography, see also Affinity chromatography; High-performance liquid chromatography (HPLC) anoxygenic phototrophic bacteria and, 219-220

antibody technologies and, 154, 158-159,166,189,192

genetically engineered microorganisms and, 78 Circular chamber, solid-state fermentation and, 112 Citric acid, anoxygenic phototrophic bacteria and, 263-264

Citrobacter freundii, hydrogen production technology and, 268 Clay minerals, genetically engineered microorganisms and, 8-9.51 Climate, anoxygenic phototrophic bacteria and, 231 Clones, antibody technologies and monoclonal antibodies, 161, 164, 170 recombinant antibodies, 172, 174, 179 -180

recombinant DNA, 191-193 Clostridium, hydrogen production technology and, 219 Clostridiurn butyricum, hydrogen production technology and, 239, 268-269, 271,279

Cocultures, anoxygenic phototrophic bacteria and, 268-269, 271, 278 Colony-forming units, genetically engineered microorganisms and, 1 1 , 1 7 Colorimetry antibody technologies and, 191 solid-state fermentation and, 130-131 Column fermentors, solid-state fermentation and, 111 Combinatorial libraries, antibody technologies and, 172-174 Competition, genetically engineered microorganisms and, 48-49 Complementarity-determining regions, antibody technologies and, 177, 179 Complete Freund’s adjuvant, antibody technologies and, 153 Compost, solid-state fermentation and, 103,109

experimental measurements, 134-135 mathematical modeling, 126-130 physical parameters, 117- 118 Concentration gradients, solid-state fermentation and, 123-128 Conversion efficiency, anoxygenic phototrophic bacteria and, 224-229 Creatine kinase, antibody technologies and, 165 Cross-linking, antibody technologies and enzyme immunoassay, 188-189 monoclonal antibodies, 155 -167 polyclonal antibodies, 151 recombinant antibodies, 177

302

INDEX

Crystallinity, solid-state fermentation and, 115,127 Crystallization, antibody technologies and, 175 Crystallography,anoxygenic phototrophic bacteria and, 256 Cyanobacteria,hydrogen production technology and, 268,280 Cyclophosphamide, antibody technologies and, 165-166 Cysteine, anoxygenic phototrophic bacteria and, 255 Cytoplasm anoxygenic phototrophic bacteria and, 251-252 solid-state fermentation and, 115 Cytotoxic T cells, antibody technologies and, 167 Cytotoxicity,antibody technologies and, 182-185

D Deamination, genetically engineered microorganisms and, 49 Degradation,solid-state fermentation and, 104-106,116,125,127 Dehydrogenases. genetically engineered microorganisms and, 3,90,94 methods of study, 16-17,36,40-41 results, 62,74,78 Deletion mutagenesis, antibody technologies and, 194 Denitrification, genetically engineered microorganisms and, 23-24,62 Dichloromethane, solid-state fermentation and, 131 2,4-Dichlorophenol,genetically engineered microorganisms and, 78 2,4-Dichlorophenoxyacetate, genetically engineered microorganisms and, 69, 74-78,90,94-95 Dielectrophoreisis, antibody technologies and, 162 Diffusion anoxygenic phototrophic bacteria and, 239-240 solid-state fermentation and, 117-116, 122.125

Diffusivity,solid-state fermentation and, 116-116,123-124,140-141 Digoxin, antibody technologies and, 177 Dinitrogen anoxygenic phototrophic bacteria and carbon assimilation, 264 classification, 217 hydrogen production, 229-230 hydrogenase, 252 nitrogenase, 240-242,244,247-249 genetically engineered microorganisms and, 3-4,46-48,94 Disulfide bonds, antibody technologies and, 174,177,180 Dithionite, anoxygenic phototrophic bacteria and, 249-250 DNA anoxygenic phototrophic bacteria and, 213,216,253,255 antibody technologies and, 151 monoclonal antibodies, 169-170 polyclonal antibodies, 154- 155 recent developments, 160 recombinant antibodies, 170,174, 176,179 recombinant DNA,191-194 genetically engineered microorganisms and, 2,4,36-37.90 solid-state fermentation and, 131 DRAG, anoxygenic phototrophic bacteria and, 245 DRAT, anoxygenic phototrophic bacteria and, 245 Drugs, antibody technologies and, 183, 185,195

E Ecologic dose, genetically engineered microorganisms and, 50,91 Ectothiorhodospim, hydrogen production technology and, 217,260 Ectothiorhodospiracae,hydrogen production technology and, 214-216 Effector cells, antibody technologies and, 181 Electron acceptors, anoxygenic phototrophic bacteria and, 257,259,263, 265-266

INDEX Electron donors, anoxygenic phototrophic bacteria and classification, 213 enzymes, 243,247 hydrogen production, 220-224, 226-227,229-230,234,236,267

Electron microscopy, solid-state fermentation and, 116 Electron paramagnetic resonance, anoxygenic phototrophic bacteria and, 256-258

Electron transport, anoxygenic phototrophic bacteria and, 242,246,250, 265

Electrophoresis, antibody technologies and, 187 ELISA, antibody technologies and, 187-189

Energy anoxygenic phototrophic bacteria and, 211-212

carbon assimilation, 259,265 enzymes, 241,243,252-253 hydrogen production, 220,233-234, 240,269

genetically engineered microorganisms and, 56 solid-state fermentation and, 105,118 Enhancers, antibody technologies and, 171 Enterobacter cloacae, genetically engineered microorganisms and, 79, 83-87

Environmental toxicants, genetically engineered microorganisms and, 50 Enzyme immunoassay, antibody technologies and, 187-191 Enzymes anoxygenic phototrophic bacteria and carbon assimilation, 260-261, 263, 266-267

hydrogen metabolism, 220 hydrogen production, 224 hydrogenase, 249-259 nitrogenase, 240-249 antibody technologies and immunofluorescence, 186 immunotherapy, 180,189-191 immunotoxins, 182-183,185 monoclonal antibodies, 166-167, 169-170

303 recombinant antibodies, 175 recombinant DNA, 191 genetically engineered microorganisms and, 3,92,94 activity, 62,68-72 metabolic activity, 8,9,12 results, 69,74 soil, 16-18, 36-34 solid-state fermentation and, 100-101 mass transfer, 105-106 mathematical modeling, 120-123, 125

physical parameters, 114-115 water activity, 112 Epitopes antibody technologies and, 151-153, 184,189,194

mapping, 156,192-194 Equilibrium, solid-state fermentation and, 112

Equilibrium relative humidity, solid-state fermentation and, 114 Equilibrium vapor pressure, solid-state fermentation and, 114 Ergosterol, solid-state fermentation and, 130-131,133

Erwinia carotovora, genetically engineered microorganisms and, 93 Escherichia coli antibody technologies and immunotoxins, 184 monoclonal antibodies, 170 polyclonal antibodies, 154 recombinant antibodies, 170-171, 175,177,179

recombinant DNA, 191 genetically engineered microorganisms and, 87 hydrogen production technology and, 239

Ethanol, anoxygenic phototrophic bacteria and, 224-226,264 Eukaryotes, antibody technologies and, 163,171-172,183

Evaporation, solid-state fermentation and, 110

EXAFS, anoxygenic phototrophic bacteria and, 257 Expression libraries, antibody technologies and, 191-192

3 04

INDEX

F

G

Fatty acids, anoxygenic phototrophic bacteria and, 216,265 Fermentation anoxygenic phototrophic bacteria and, 219,263-266,268-269 antibody technologies and, 164,194 genetically engineered microorganisms and, 35 solid-state, see Solid-state fermentation submerged, see Submerged fermentation Fibrin, antibody technologies and, 167 Fick’s law, solid-state fermentation and, 117,127 Flow cytometry, antibody technologies and, 187 Fluidized bed reactor, anoxygenic phototrophic bacteria and, 237 Fluorescein isothiocyanate (FITC),antibody technologies and, 167,187 Fluorescence antibody technologies and, 186-187, 191 solid-state fermentation and, 131 Fluorescence activated cell sorter,antibody technologies and, 167 Fluorimetry, solid-state fermentation and, 131-132 Forced air, solid-state fermentation and,

Gas anoxygenic phototrophic bacteria and, 239,247-249,279 solid-state fermentation and, 101 experimental measurements, 134-140,142 mass transfer, 102-103 mathematical modeling, 118,126,130 physical parameters, 117 Gas chromatography, anoxygenic phototrophic bacteria and, 219-220 Gas chromatography-mass spectroscopy, genetically engineered microorganisms and, 78 Gene expression, antibody technologies and, 170-172 Gene transfer, genetically engineered microorganisms and, 51,91 Genes, anoxygenic phototrophic bacteria and, 251,254-255 Genetic engineering, antibody technologies and, 156,172,184-185 Genetic markers, antibody technologies and, 167 Genetically engineered microorganisms, 2-7.90-95 methods of study dinitrogen, 46-48 growth rates, 48-49 metabolic activity, 8-16 microbial assays, 18-36 nitrogen transformations, 41-46 soil enzymes, 36-41 soil preparation, 7-8, 16-18 statistics, 49-50 results, 50-52 2,4-dichlorophenoxyacetate,69, 74-78 metabolic activity, 52-55 nitrogen, 78-87 pH, 69,73 soil enzymes, 62,68-72 species diversity, 53,56-67 survival, 87-90 Genotype, genetically engineered microorganisms and, 11 Glucosamine, solid-state fermentation and, 120,130-133

108

Forced convection, solid-state fermentation and, 112 Formate, anoxygenic phototrophic bacteria and, 220,261-262,266,269 Fructose, anoxygenic phototrophic bacteria and, 264, 266 Fungal propagules, genetically engineered microorganisms and, 21,56,78,90, 94 Fungi, solid-state fermentation and, 100-101,112 experimental measurements, 130- 131, 134 physical parameters, 115

305

INDEX Glucose anoxygenic phototrophic bacteria and carbon assimilation, 264 hydrogen production, 220,224-226, 228,268-269

genetically engineered microorganisms and metabolic activity, 52-53 results, 74,87,90 soil enzymes, 62,69 species diversity, 56,62 solid-state fermentation and, 121-123 Glutamate, anoxygenic phototrophic bacteria and, 223,225,230,244,247 Glutamate dehydrogenase, anoxygenic phototrophic bacteria and, 244 Glutamine, anoxygenic phototrophic bacteria and, 244,246-247. 264 Glutamine synthetase, anoxygenic phototrophic bacteria and, 244,246 Glutaraldehyde, antibody technologies and, 151,165,188-189 Glycoprotein, antibody technologies and, 188,193-194

Glycosylation, antibody technologies and, 171

GOGAT, anoxygenic phototrophic bacteria and, 244,246 Green bacteria, hydrogen production technology and, 215-217,220,260 Growth factors antibody technologies and, 163 solid-state fermentation and, 100 Growth factors medium, genetically engineered microorganisms and, 29-30

Heterochimeras, antibody technologies and, 174-175 Heterolysis, anoxygenic phototrophic bacteria and, 258-259 High-performance liquid chromatography (HPLCl antibody technologies and, 158-160 genetically engineered microorganisms and, 78 solid-state fermentation and, 131 HIV-1,antibody technologies and, 174-176,188,193-194

Homochimeras, antibody technologies and, 175-177 Homology anoxygenic phototrophic bacteria and, 253,255

antibody technologies and, 152 genetically engineered microorganisms and, 4-5,91-92,94 metabolic activity, 9,11, 52-53 methods of study, 35-36,48, 50 nitrogen transformations, 78-79,87 results, 51-52, 69,90 soil preparation, 18 species diversity, 53,56,62 Homolysis, anoxygenic phototrophic bacteria and, 258 Hormones, antibody technologies and, 152

Horseradish peroxidase, antibody technologies and, 189-191 Humidity genetically engineered microorganisms and, 18 solid-state fermentation and, 108,110, 112-114

H Hapten, antibody technologies and, 169-170

Heat, solid-state fermentation and, 113, 118

dissipation, 111-112 transfer, 106-108, 142 experimental measurements, 135, 140-141

mathematical modeling, 118,126,128 Hepatoma cells, antibody technologies and, 155

Humoral response, antibody technologies and, 154 Hybridomas, antibody technologies and, 161-167,175

Hybrids anoxygenic phototrophic bacteria and, 213

antibody technologies and, 166,172, 174

Hydride, anoxygenic phototrophic bacteria and, 258-259 Hydrochloric acid, genetically engineered microorganisms and, 10, 12,15-16

306

INDEX

Hydrogen production technology, anoxygenic phototrophic bacteria and, see Anoxygenic phototrophic bacteria Hydrogenase, anoxygenic phototrophic bacteria and, 241-243, 249-259 carbon assimilation, 261 classification, 218 hydrogen metabolism, 219-220 hydrogen production, 227-228, 231, 239-240

Hydrophobic interaction chromatography, antibody technologies and, 159

enzyme immunoassay, 190 immunoglobulin-binding proteins, 157-158

immunotherapy, 180-182 immunotoxins, 183,185 monoclonal antibodies, 163-167 polyclonal antibodies, 154-155 radiolabels, 186 recent developments, 158,160-161 recombinant antibodies, 170-172, 174-175,177-178

Immunology, antibody technologies and, 150,156

I Idiotypes, antibody technologies and, 182, 194

polyclonal antibodies, 155-157 recombinant antibodies, 172,174 Illumination, anoxygenic phototrophic bacteria and, 271-273, 279 Imidazole, antibody technologies and, 169 Immobilization supports, antibody technologies and, 187-191 Immobilized cells, anoxygenic phototrophic bacteria and enzymes, 249 hydrogen production, 224, 227, 234-240

hydrogen production technology, 268, 270,273-278

Immune responses, antibody technologies and, 151-154,162,173,175 Immunization, antibody technologies and, 156

immunotherapy, 182 monoclonal antibodies, 164-166, 168 polyclonal antibodies, 152, 157 Immunoassays, antibody technologies and, 151,186,194 enzyme immunoassay, 187-191 immunoglobulin-binding proteins, 158 immunotherapy, 180 polyclonal antibodies, 152 recombinant DNA, 191 Immunogenicity, antibody technologies and, 152-153,165,177,180-81 Immunoglobulin, antibody technologies and, 151,194-195

Immunoreactivity, antibody technologies and, 160 Immunoscreening, antibody technologies and, 191-193 Immunosuppression, antibody technologies and, 195 Immunotherapy, antibody technologies and, 151,180-182 polyclonal antibodies, 156-157 radiolabels, 186 recombinant antibodies, 175,177 Immunotoxins, antibody technologies and, 151,182-185,194 Mammatory response, antibody technologies and, 153 Infrared analysis, solid-state fermentation and, 133-135 Inhibitors anoxygenic phototrophic bacteria and carbon assimilation, 260,263 classification, 219 enzymes, 241-242,245-247 hydrogen production, 230,237,240, 271

antibody technologies and, 155, 167, 169,181,184,195

genetically engineered microorganisms and, 74,78,90,92 solid-state fermentation and, 114, 125 Insulin, antibody technologies and, 155, 163

Interferon, antibody technologies and, 167

Interleukin-2, antibody technologies and, 175,177,184-185

Interparticle mass transfer, solid-state fermentation and, 103-104

307

INDEX Intraparticle mass transfer, solid-state fermentation and, 104 Iodine, antibody technologies and, 186 Ion-exchange chromatography, antibody technologies and, 159 Iron, anoxygenic phototrophic bacteria and hydrogenase, 253,255-258 nitrogenase, 243,245,247-248

Ligands anoxygenic phototrophic bacteria and, 253,257-258

antibody technologies and, 155,160 Light anoxygenic phototrophic bacteria and, 212

advances, 262,264 carbon assimilation, 259,262, 264-266

K

classification, 216-217 hydrogen production, 220, 229, 231-234,238,240

Kanamycin, genetically engineered microorganisms and, 93 Kaolinite, genetically engineered microorganisms and, 51 Kinetics anoxygenic phototrophic bacteria and, 212,220,256,266

antibody technologies and, 168 genetically engineered microorganisms and, 9,41 solid-state fermentation and, 101-102, 104,142

mathematical modeling, 118-123, 125

physical parameters, 114,117 Klebsiella, anoxygenic phototrophic bacteria and, 247 Klebsiella pneumoniae, anoxygenic phototrophic bacteria and, 268,271 Koji room, solid-state fermentation and, 108,120,128

L

hydrogenase, 257 nitrogenase, 241444,249 genetically engineered microorganisms and, 5 Light scattering, solid-state fermentation and, 132-133 Lignin peroxidase, genetically engineered microorganisms and, 93 Lipids anoxygenic phototrophic bacteria and, 213,216

antibody technologies and, 153. 155 Lipopolysaccharides anoxygenic phototrophic bacteria and, 213

antibody technologies and, 154 Liposomes, antibody technologies and, 182

Lymphocytes, antibody technologies and immunotherapy, 180 monoclonal antibodies, 163,165 panning, 165,167 recombinant antibodies, 172-173 Lymphoma, antibody technologies and, 182,187

Lactate, anoxygenic phototrophic bacteria and enzymes, 247 hydrogen production, 223-228,231, 233,236-237

hydrogen production technology, 269, 275

Lactose, antibody technologies and, 184 Lectin, genetically engineered microorganisms and, 93 Leukemia, antibody technologies and, 182.187

M Maintenance medium, genetically engineered microorganisms and, 18-19 Malate, anoxygenic phototrophic bacteria and carbon assimilation, 260-261, 266 enzymes, 236,238-240,247 hydrogen production, 223-230.236, 238-240

308

INDEX

Malignancy, antibody technologies and, 185

Mass diffusivities, solid-state fermentation and, 116-117 Mass spectrometry, anoxygenic phototrophic bacteria and, 220 Mass transfer, solid-state fermentation and, 102-106 bioreactor design, 108-111 experimental measurements, 140- 141 mathematical modeling, 118,124-126 physical parameters, 116- 117 Mass transport, solid-state fermentation and, 202,135,142 Matric potential, solid-state fermentation and, 113 MBS, antibody technologies and, 152 Memory cells, antibody technologies and, 195

Messenger RNA, antibody technologies and, 155 Metabolic activity, genetically engineered microorganisms and methods of study, 8-16 results, 52-55, 62,74 Metabolism, solid-state fermentation and, 101,106-107,112-113

Methane, anoxygenic phototrophic bacteria and, 280-281 Methanol, anoxygenic phototrophic bacteria and, 263-264 Microbial assays, genetically engineered microorganisms and, 16-36 Microbial cells, Solid-state fermentation and, 100,102,132 Microbial growth, solid-state fermentation and experimental measurements, 135 mathematical modeling, 118,125-126, 129

physical parameters, 116-1 17 water activity, 112 Mineralization, genetically engineered microorganisms and, 36,41,53,90,94 Minimum inhibitory concentration, genetically engineered microorganisms and, 35 Moisture genetically engineered microorganisms and, 8

solid-state fermentation and, 103,107, 111-113

Molybdenum, anoxygenic phototrophic bacteria and, 248,263 Monoclonal antibodies, technologies, 150-152,161-170

enzyme immunoassay, 191 immunofluorescence, 187 immunotherapy, 180-181 radiolabels, 186 recent developments, 159-160 recombinant antibodies, 172-173, 175, 177

recombinant DNA, 191-192,194 Montmorillonite, genetically engineered microorganisms and, 51,79 Multiple antigenic peptides, antibody technologies and, 152 Muramyl dipeptide, antibody technologies and, 153-154 Muscle creatine kinase, antibody technologies and, 165 Mutagenesis, antibody technologies and, 173-174,179

Mutants anoxygenic phototrophic bacteria and, 228-229,242, 246,252,266

antibody technologies and, 184 genetically engineered microorganisms and, 35,93 Myasthenia gravis, antibody technologies and, 156 Mycelium, solid-state fermentation and bioreactor design, 109-110 experimental measurements, 132 mass transfer, 104 mathematical modeling, 119,122-123 water activity, 113 Mycobacteria, antibody technologies and, 153,154

Myeloma cells, antibody technologies and, 161-162,164,171 Myosin, antibody technologies and, 192

N Neisserio meningitidis, antibody technologies and, 154 Neomycin, antibody technologies and, 166

INDEX Neutralization, genetically engineered microorganisms and, 16 Nickel, anoxygenic phototrophic bacteria and, 253,255-258,263 nif genes anoxygenic phototrophic bacteria and, 247

genetically engineered microorganisms and, 4,44 Nitrate anoxygenic phototrophic bacteria and,

309

metabolic activity, 8-9 methods of study, 41,48,50 microbial assays, 35-36 results, 69 n-propanol, anoxygenic phototrophic bacteria and, 223-226,228,264 Nucleases, genetically engineered microorganisms and, 37 Nucleotides, antibody technologies and, 179

219

genetically engineered microorganisms and, 62,87 Nitrate broth, genetically engineered microorganisms and, 23-24 Nitrification, genetically engineered microorganisms and, 4 methods of study, 9,49 microbial assays, 24-26 nitrogen transformations, 41-46, 79,87 results, 79,87 Nitrite, genetically engineered microorganisms and, 62,87 Nitrocellulose, antibody technologies and,

0

Oligonucleotides anoxygenic phototrophic bacteria and, 213

antibody technologies and, 179 Optical absorption spectroscopy, anoxygenic phototrophic bacteria and, 256 Organic acids, anoxygenic phototrophic bacteria and, 221,227,261,264, 268-269

Osmotic potential, solid-state fermentation and, 113 187-188,191 Osmotic pressure, solid-state fermentation Nitrogen and, 118 anoxygenic phototrophic bacteria and Osmotic stress, anoxygenic phototrophic carbon assimilation, 259,264,267 bacteria and, 240,251 classification, 216-217, 219 Outer membrane proteins, antibody techenzymes, 241-242,244,246-249,252 nologies and, 154-155 hydrogen production, 223,227,230, Oxidation 267,271 anoxygenic phototrophic bacteria and, genetically engineered microorganisms 249,263 and, 3,94 antibody technologies and, 166 dinitrogen fixation, 46-48 genetically engineered microorganisms metabolic activity, 11 and, 36 transformations, 41-46, 78-87 solid-state fermentation and, 126-127 solid-state fermentation and, 100,123, Oxidizers, genetically engineered micro131,133 organisms and, 62 Nitrogenase, anoxygenic phototrophic Oxygen bacteria and, 240-249, 252-253 anoxygenic phototrophic bacteria and classification, 218 carbon assimilation, 259,263 hydrogen metabolism, 219 enzymes, 242,247,252,257 hydrogen production, 228,231, hydrogen production, 240 239-240,278 solid-state fermentation and, 100-101 Nodulation, genetically engineered microbioreactor design, 108-110 organisms and, 93 experimental measurements, 132, Novel genes, genetically engineered mi134,140 croorganisms and, 2-3, 5,90-94 heat transfer, 106-107

310

INDEX

mass transfer, 102-105 mathematical modeling, 118,120, 122,124-127

physical parameters, 114,117-118 Oxygen diffusion, solid-state fermentation and. 104-106,109

P Packed bed fermentor, solid-state fermentation and bioreactor design, 108-110 experimental measurements, 135-1 37 heat dissipation, 112 mathematical modeling, 128-129 Pelochromatium,hydrogen production technology and, 216 Peptides, antibody technologies and enzyme immunoassay, 187-189 immunotoxins, 183-184 monoclonal antibodies, 169 polyclonal antibodies, 151-153 recombinant antibodies, 177,179-180 recombinant DNA, 192 Percolation, solid-state fermentation and, 110

Perfusion, genetically engineered microorganisms and, 3,94 methods of study, 49 nitrogen transformations, 41-43, 78 Peripheral blood lymphocytes, antibody technologies and, 163,167 PH anoxygenic phototrophic bacteria and carbon assimilation, 263,266 classification, 217 enzymes, 243,253 hydrogen production, 227,229-231, 240

hydrogen production technology, 272-276,279

antibody technologies and, 160 genetically engineered microorganisms and, 4 methods of study, 12,17,41 results, 69,73,78-79 solid-state fermentation and, 101,118, 135

Phenotype antibody technologies and, 187,193 genetically engineered microorganisms and, 90-91,95 methods of study, 11,32-36 results, 52,90 Phospholipids, antibody technologies and, 155 Phosphomonoesterase,genetically engineered microorganisms and, 37-38 Phosphorylation anoxygenic phototrophic bacteria and, 265

solid-state fermentation and, 103 Photobioreactors, anoxygenic phototrophic bacteria and, 249,270-279 Photosynthesis, anoxygenic phototrophic bacteria and, 212,279-281 carbon assimilation, 261,266 classification, 213, 216 enzymes, 250-254,257 hydrogen production, 220,229,234, 268-269

Phototrophic bacteria, anoxygenic, see Anoxygenic phototrophic bacteria Physiological groups, genetically engineered microorganisms and, 28-32 Plasma membrane, antibody technologies and, 155 Plasmids, genetically engineered microorganisms and, 4, 79,90,93 Plasmodium falciparum, antibody technologies and, 189 p-nitrophenyl, genetically engineered microorganisms and, 36 Polyclonal antibodies, technologies, 150-157

immunotoxins, 182 recent developments, 160 recombinant DNA,191-192 Polymerase chain reaction, antibody technologies and, 172-173 Polypeptides anoxygenic phototrophic bacteria and, 245

antibody technologies and enzyme immunoassay, 187 immunotoxins, 183 polyclonal antibodies, 155

INDEX recombinant antibodies, 171, 174-175,177

recombinant DNA,192 Polysaccharides, anoxygenic phototrophic bacteria and, 268 Polyvinyl difluoride (PVDF),antibody technologies and, 188 Population dynamics, genetically engineered microorganisms and, 10 Porosity, solid-state fermentation and, 114,116,118

Potassium, genetically engineered microorganisms and, 36 Potassium hydroxide, genetically engineered microorganisms and, 11 - 14 Precipitation, antibody technologies and, 158

Prokaryotes anoxygenic phototrophic bacteria and, 213

antibody technologies and, 170-171 Promoters, antibody technologies and, 171

Proteases, genetically engineered microorganisms and, 37 Protein anoxygenic phototrophic bacteria and, 279-280

carbon assimilation, 266 hydrogen production technology, 268 hydrogenase, 250,253-257 nitrogenase, 243,245,247-248 antibody technologies and, 151,194 enzyme immunoassay, 187-189,191 immunoglobulin-binding proteins,

311

Protoplasts antibody technologies and, 171 genetically engineered microorganisms and, 93 Protozoa, genetically engineered microorganisms and, 26-28 Pseudomonas, hydrogen production technology and, 261 Pseudomonas exotoxin, antibody technologies and, 184-185 Pseudomonas jluorescens, hydrogen production technology and, 268-269 Pseudomonas putida, genetically engineered microorganisms and, 90,94 microbial assays, 20-21 results, 69,74,87 Purification anoxygenic phototrophic bacteria and, 267,281

antibody technologies and, 151, 158-161

immunotherapy, 181 monoclonal antibodies, 163 recombinant antibodies, 175,180 recombinant DNA,192 Purple bacteria, hydrogen production technology and, 214-216,220,251, 259-260

Purple nonsulfur bacteria, hydrogen production technology and, 213, 216-217,223,259-260

Pyruvate, anoxygenic phototrophic bacteria and, 223,225-226,266

157-158

immunotherapy, 181 immunotoxins, 183-185 monoclonal antibodies, 163,165 polyclonal antibodies, 154-156 radiolabels, 186 recent developments, 159-160 recombinant antibodies, 174,176-177 recombinant DNA, 191-194 solid-state fermentation and, 115 Proteolysis, antibody technologies and, 152,169,177,180,183,194

Proteosomes, antibody technologies and, 154

Q Quil A,antibody technologies and, 154 Quillaja saponaria, antibody technologies and, 154

R Radioactivity, antibody technologies and, 186

Radioimmunoassays, antibody technologies and, 186

312

INDEX

Radioimmunodetection, antibody technologies and, 186 Radioimmunotherapy, antibody technologies and, 186 Radiolabeled antibody technologies, 186 Radionuclides, antibody technologies and, 186

Rc2, anoxygenic phototrophic bacteria and, 245-246 Reactors, solid-state fermentation and, 107 Recombinant antibodies, technologies, 150-151,170-180,184,194

Recombinant DNA,antibody technologies and, 191-194 Recombinant proteins, antibody technologies and, 157-158,170,192 Replication, genetically engineered microorganisms and, 3 metabolic activity, 10-12 results, 54,62, 74 Resistance, genetically engineered microorganisms and, 9 1 Respiration anoxygenic phototrophic bacteria and, 247,252

solid-state fermentation and, 107,119, 124-125

Rhizopus oligosporus, solid-state fermentation and, 105-106,121-122 Rhodobacillus pdustris, hydrogen production technology and, 220,228, 234,236-239

advances, 267 carbon assimilation, 265-266 enzymes, 243-244,248-249 Rhodobacter. hydrogen production technology and, 216-217,233 Rhodobacter capsulatus, hydrogen production technology and, 223,

carbon assimilation, 261, 264 classification, 216 enzymes, 244,246 Rhodobacter sulfidophilus, hydrogen production technology and enzymes, 243-244,253 hydrogen production, 221, 229,231, 270

Rhodocyclus purpureus, hydrogen production technology and, 240, 244 Rhodopseudomonas, hydrogen production technology and, 220,223, 227-228,235-236,238,

240

advances, 269 -27 1 carbon assimilation, 261,265 enzymes, 247 Rhodopseudomonas acidophila, hydrogen production technology and, 217, 263

Rhodospirillum rubrum, hydrogen production technology and, 220, 224, 229-230,232-240

advances, 268-270, 275-278 carbon assimilation, 261,263,265-266 h ydrogenase , 25 1-252 nitrogenase, 244-248 Ribosomes, antibody technologies and, 174,185

Ribulose 1,5-bisphosphate carboxylase, anoxygenic phototrophic bacteria and, 259-260 Ricin, antibody technologies and, 183-184

Rifampin, genetically engineered microorganisms and, 93 Rocking drum reactor, solid-state fermentation and, 110-111 Rotating drum fermentor, solid-state fermentation and, 108-111

227-233,235-236,239

advances, 268-270,272 carbon assimilation, 261, 264-265 classification, 216 hydrogen metabolism, 220 hydrogenase, 251-255 nitrogenase. 241,243,245-248 Rhodobacter sphaeroides, hydrogen production technology and, 227-232, 234-236,239

advances, 269-270,274,276

S Saccharomyces cerevisiae, antibody technologies and, 1 7 1 Saponin, antibody technologies and, 154, 185 SDS-PAGE,antibody technologies and, 187-188,192

INDEX Secondary metabolites, solid-state fermentation and, 100 Selective markers, genetically engineered microorganisms and, 92 Sequences anoxygenic phototrophic bacteria and, 216,255

antibody technologies and enzyme immunoassay, 189 monoclonal antibodies, 165,169-170 recombinant antibodies, 172,174,177 recombinant DNA, 192 Serum-free medium, antibody technologies and, 163-164 Single cell protein, anoxygenic phototrophic bacteria and, 268,279-280 Single-chain antibodies, antibody technologies and, 177-179 Site-directed mutagenesis, antibody technologies and, 174 Sodium hydroxide genetically engineered microorganisms and, 9-12,14-16 solid-state fermentation and, 132 Soil, genetically engineered microorganisms and, 4-7,90-95 2,4-dichlorophenoxyacetate,69,74- 78 enzymes, 36-41,62,68-72 growth rates, 48 metabolic activity, 8-9,11,15-16, 52-53

methods of study, 49 microbial assays, 16-18, 32-36 nitrogen transformations, 41-43, 78,87 PH, 69 preparation, 7-8 results, 50-52 species diversity, 53,56,62 survival, 87-90 Soil extract agar, genetically engineered microorganisms and, 19-20 Soil extract medium, genetically engineered microorganisms and, 30-31, 36

Solid-state fermentation, 99- 102, 141-142

bioreactor design, 108-111 experimental measurements biomass, 130-134 diffusivity, 140-141

313

temperature, 134-140 heat dissipation, 111-112 heat transfer, 106-108 mass transfer, 102-103 degradation, 105-106 interparticle, 103-104 intraparticle, 104 oxygen diffusion, 104-105 mathematical modeling, 118 concentration gradients, 123-128 kinetics, 119-123 temperature gradients, 128-130 nomenclature, 142-144 physical parameters, 114-118 water activity, 112-114 Soybean, genetically engineered microorganisms and, 93 Species diversity, genetically engineered microorganisms and, 92,94 nitrogen transformations, 41 results, 53,56-67 Spectroscopy anoxygenic phototrophic bacteria and, 256-258

genetically engineered microorganisms and, 78 Staphylococcal protein A, antibody technologies and, 157-158,160 Streptavidin, antibody technologies and, 162

Streptococcal protein G,antibody technologies and, 157-158 Streptornyces Jividans, genetically engineered microorganisms and, 93 Subclones, antibody technologies and, 193-194

Submerged fermentation, 100-101, 134, 141

bioreactor design, 108 heat transfer, 106 mass transfer, 102,104-105 mathematical modeling, 123-125 Substrate conversion efficiency, anoxygenic phototrophic bacteria and, 224-229

Succinate, anoxygenic phototrophic bacteria and, 224-226, 260,266 Sucrose anoxygenic phototrophic bacteria and, 226

314

INDEX

solid-state fermentation and, 133 sugar anoxygenic phototrophic bacteria and, 265-266

solid-state fermentation and, 131 Sulfatases, genetically engineered microorganisms and, 17 , 7 8 Sulfur anoxygenic phototrophic bacteria and carbon assimilation, 260 classification,213,217 enzymes, 243,253,255-258 hydrogen production, 220 genetically engineered microorganisms and, 3 , 3 6 Surrogate receptors, antibody technologies and, 155 Synechococcus cedrorum, hydrogen production technology and, 268-269 Syntex adjuvant formulation, antibody technologies and, 153 Synthetic peptides, antibody technologies and, 151-153

Thermal diffusivity, solid-state fermentation and, 117-118,140-141 Thermoactinomyces, solid-state fermentation and, 116 Thermodynamic equilibrium, solid-state fermentation and, 114 Thermodynamics, solid-state fermentation and, 118 Thiocapsa roseopersicina, hydrogen production technology and, 214, 250-254,256-257

Tobacco, antibody technologies and, 171 Toxicants, genetically engineered microorganisms and, 50 Toxicity anoxygenic phototrophic bacteria and, 280

antibody technologies and, 153-154 genetically engineered microorganisms and, 74 Toxins, antibody technologies and, 182-183

Transcription, antibody technologies and, 155,171

Transfection, antibody technologies and,

T T cell receptor, antibody technologies and, 172,174 T cells, antibody technologies and, 194-195

immunotherapy, 181 polyclonal antibodies, 152-153 radiolabels, 186 recombinant antibodies, 176 Temperature anoxygenic phototrophic bacteria and, 227,229,231,256,272-273

genetically engineered microorganisms and, 5,8-9, 18 solid-state fermentation and, 101, 142 experimental measurements, 131, 141 heat dissipation, 111-112 heat transfer, 106-108 mathematical modeling, 118, 127-130

physical parameters, 117-1 18 water activity, 112-113 Thermal conductivity, solid-state fermentation and, 129-130,140

171

Transfectomas,antibody technologies and, 172 Transferrin, antibody technologies and, 16 3, 184,191

Translation, antibody technologies and, 155,171

Transplantation, antibody technologies and, 186 Transplantation immunotherapy, antibody technologies and, 181,195 Tray fermentor, solid-state fermentation and, 108,111,135-136.138-140 Trickling filter reactor, solid-state fermentation and, 110 Triphenylformazan, genetically engineered microorganisms and, 36 Trypanosoma, antibody technologies and, 156

Tumor necrosis factor, antibody technologies and, 167 Tumors, antibody technologies and, 177, 180-182,184-186

Tyrosine kinase, antibody technologies and, 175

315

INDEX

V Vaccines, antibody technologies and, 152-153,156-157,194-195

Vacuoles, antibody technologies and, 189 Velocity, solid-state fermentation and, 112 Viscosity, anoxygenic phototrophic bacteria and, 239 Vitamin B,, medium, genetically engineered microorganisms and, 31-32 Vitamins, anoxygenic phototrophic bacteria and, 217,280

Water tension, genetically engineered microorganisms and, 9,11-12 Water vapor, solid-state fermentation and, 107

Wheat, solid-state fermentation and, 101 X

X-ray crystallography,antibody technologies and, 156,179 Xenobiotics, genetically engineered microorganisms and, 74

W Y Wastewater utilization, anoxygenic phototrophic bacteria and, 267-268 Water activity, solid-state fermentation and, 112-114,142 Water content, solid-state fermentation and, 108 Water potential, solid-state fermentation and, 112-113

Yeast anoxygenic phototrophic bacteria and, 230

antibody technologies and, 170-171 solid-state fermentation and, 100 Yeast extract medium, genetically engineered microorganisms and, 30-32

CONTENTS OF PREVIOUS VOLUMES Volume 28 Immobilized Plant Cells P. Brodelius and K. Mosbach Genetics and Biochemistry of Secondary Metabolism Vedpal Singh Malik Partition Affinity Ligand Assay [PALA): Applications in the Analysis of Haptens, Macromolecules, and Cells Bo Mattiasson, Matts Ramstorp, and Torbjorn G. 1. Ling Accumulation, Metabolism, and Effects of Organophosphorus Insecticides on Microorganisms Rup La1 Solid Substrate Fermentations K. E. Aidoo, A. Hendry, and B. J. B. Wood Microbiology and Biochemistry of Miso [Soy Paste] Fermentation Sumbo H. Abiose, M. C. Allan, and B. J. B. Wood INDEX

New Perspective on Aflatoxin Biosynthesis J. W. Bennett and Siegfried B. Christensen Biofilms and Microbial Fouling W. G. Chamcklis and K. E. Cooksey Microbial Influences: Fermentation Process, Properties, and Applications Erick J. Vandamme and Dirk G. Derycke Enumeration of Indicator Bacteria Exposed to Chlorine Gordon A. McFeters and Anne K. Camper Toxicity of Nickel to Microbes: Environmental Aspects H. Babich and G . Stotzky INDEX

Volume 30 Interactions of Bacteriophages with Lactic Streptococci Todd B. Klaenhammer

Volume 29

Microbial Metabolism of Polycyclic Aromatic-Hydrocarbons Carl E. Cerniglia

Stabilization of Enzymes against Thermal Inactivation Alexander M. Klibanov

Microbiology of Potable Water Betty H. Olson and Laslo A. Nagy

Production of Flavor Compounds by Microorganisms G. M. Kempler

Applied and Theoretial Aspects of Virus Adsorption to Surfaces Charles P. Gerba 316

CONTENTS OF PREVIOUS VOLUMES Computer Applications in Applied Genetic Engineering Joseph L. Modelevsky Reduction of Fading of Fluorescent Reaction Product for Microphotometric Quantitation G. L. Picciolo and D. S. Kaplan

317

Problems and Alternative Approaches to Identification N. Robert Ward, Roy L. Wolfe, Carol A. Justice, and Betty H. Olson INDEX

Volume 32 INDEX

Microbial Corrosion of Metals Warren P. Iverson Volume 31 Genetics and Biochemistry of Clostridium Relevant to Development of Fermentation processes Palmer Rogers The Acetone Butanol Fermentation B. McNeil and B. Kristiansen Survival of, and Genetic Transfer by, Genetically Engineered Bacteria in Natural Environments G. Stotzky and H. Babich Apparatus and Methodology for Microcarrier Cell Culture S.Reuveny and R. W. Thoma Naturally Occurring Monobactams William L. Parker, Joseph O’Sullivan, and Richard B. Sykes

Economics of the Bioconversion of Biomass to Methane and Other Vendable Products Rudy J. Wodzinski, Robert N . Gennaro, and Michael H. Scholla The Microbial Production of 2,3Butanediol Robert J, Magee and Nain Kosaric Microbial Sucrose Phosphorylase: Fermentation Process, Properties, and Biotechnical Applications Erick J. Vandamme, Jan Van Loo, Lieve Machtelinckx, and Andre De Loports Antitumor Anthracyclines Produced by Streptomyces peucetius A. Grein INDEX

New Frontiers in Applied Sediment Microbiology Douglas Gunnison

Ecology and Metabolism of Thermomatrix thiopara Daniel K. Brannan and Douglas E. Caldwell Enzyme-Linked Immunoassays for the Detection of Microbial Antigens and Their Antibodies John E. Herrmann The Identification of Gram-Negative, Nonfermentative Bacteria from Water:

Volume 33 The Cellulosome of Clostridium thermocellum Raphael Lamed and Edward A. Bayer Clonal Populations with Special Reference to Bacillus sphaericus Samuel Singer Molecular Mechanisms of Viral Inactivation by Water Disinfectants A. B. Thurman and C. P. Gerba

318

CONTENTS OF PREVIOUS VOLUMES

Microbial Ecology of the Terrestrial Subsurface William C. Ghiorse and John T. Wilson Foam Control in Submerged Fermentation: State of the Art N. P. Ghildyal, B. K. Lonsane, and N. G. Karanth Applications and Mode of Action of Formaldehyde Condensate Biocides H. W. Rossmoore and M. Sondossi Occurrence and Mechanisms of Microbial Oxidation of Manganese Kenneth H. Nealson, Bradley M. Tebo, and Reinhardt A. Rosson Recovery of Bioproducts in China: A General View Xiong Zhenping

Volunn 36 Production of Bacterial Thermostable aAmylase by Solid-state Fermentation: A Potential Tool for Achieving Economy in Enzyme Production and Starch Hydrolysis B. K. Lonsane and M. V. Ramesh Methods for Studying Bacterial Gene Transfer in Soil by Conjugation and Transduction G. Stotzky, Monica A. Devanas, and Lawrence A. Zeph Microbial Levan Youn W.Han Review and Evaluation of the Effects of Xenobiotic Chemicals on Microorganisms in Soil R. J. Hicks, G. Stotzky, and P. Van Voris

INDEX

Volume 34 What’s in a Name?-Microbial Secondary Metabolism J. W. Bennett and Ronald Bentley Microbial Production of Gibberellins: State of the Art P. K. R. Kumor and B. K. Lonsane Microbial Dehydrogenations of Monosaccharides MiloS KulhCInek Antitumor and Antiviral Substances from Fungi Shung-Chang Jong and Richard Donovick Biotechnology-The Golden Age V. S . Malik INDEX

Disclosure Requirements for Biological Materials in Patent Law Shung-Chong Jong and Jeannette M. Birmingham INDEX

Volume 36 Microbial Transformations of Herbicides and Pesticides Douglas J. Cork and James P. Krueger An Environmental Assessment of Biotechnological Processes M. S. Thakur, M. J. Kennedy, and N. G. Karanth Fate of Recombinant Escherichia coli K-12Strains in the Environment Gregg Bogosian and James F. Kane Microbial Cyctochromes P-450and Xenobiotic Metabolism F. Sima Sariaslani

CONTENTS OF PREVIOUS VOLUMES Foodborne Yeasts T.Dedk High-Resolution Electrophoretic Purification and Structural Microanalysis of Peptides and Proteins Erik P. Lillehoj and Vedpal S. Malik

319

Haloperoxidases: Their Properties and Their Use in Organic Synthesis M. C. A. Franssen and H. C. van der Plas Medicinal Benefits of the Mushroom Ganoderma S. C. Jong and J. M. Birmingham

INDEX

Volume 37

Microbial Degradation of Biphenyl and Its Derivatives Frank K. Higson

Microbial Degradation of the Nitroaromatic Compounds Frank K. Higson

The Sensitivities of Biocatalysts to Hydrodynamic Shear Stress Ales Prokop and Rakesh K. Bajpai

An Evaluation of Bacterial Standards and

Biopotentialities of the Basidiomacromycetes Somasundaram Rajarathnam, Mysore Nanjarajurs Shashirekha, and Zakia Bano

Disinfection Practices Used for the Assessment and Treatment of Stormwater Marie L. O’Shea and Richard Field

INDEX



ISBN 0-12-002638-4

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