RICE UNIVERSITY
ISOLATION, CHARACTERIZATION AND SUBSTRATE-TRANSPORT STUDIES OF A NEW, UNIQUE METHYLOTROPH
by
Thomas Alan Keuer
A THESIS SUBMITTED IN PARTIAL FULFULLMENT OF THE REQUIREMENTS FOR THE DEGREE
Master of Science
APPROVED, THESIS COMMITTEE:
E. Terry Papoutsakis, Assistant Professor of Chemical Engineering
LarryOf. Mclntire, Professor and Chairman of Chemical Engineering
<03^ ' Roger Storck, Professor of Biology
Houston, Texas
April, 1984
3 1272 00289 0232 ABSTRACT
Keuer, Thomas A. M.S. Rice University, April 1984. Isolation, Characterization and Substrate-Transport Studies of a New, Unique Methylotroph. Major Professor: E. T. Papoutsakis.
Methylotrophic bacteria which assimilate carbon via the Ribulose Monophosphate Pathway are bioenergetically superior to other methylotrophs. The dehydrogenases which catalyze the oxidation of formaldehyde to formate and formate to CO2 in RMP bacteria produce much of the
ATP required for biosynthesis. A strain, designated T15, has been isolated on the basis of high In vitro activities of the above two key enzymes, and has been biochemically characterized. The new strain exhibits high yields (up to 0.63 g cells/g MeOH) and growth rates (up to 0.46 hr“^) in batch culture? however, the yields and growth rates in continuous culture are significantly lower.
Study of the transport mechanisms has provided valuable insight into the relationship between substrate uptake and the growth characteristics of T15. Experi¬ ments with radiolabelled substrates have indicated that methanol enters the cells primarily by diffusion? consequently, the bacteria are not able to accumulate methanol internally in order to support efficient Ill
continuous growth. Formaldehyde, on the other hand, is accumulated by an active transport system which depends on the A pH component of the membrane proton-motive force.
The formate uptake mechanism is also dependent on the
ApH, but ‘is more complex, possibly due to the uncoupling effect of the organic acid on the cell membrane. ACKNOWLEDGMENTS
I would like to recognize those whose assistance
made the completion of this thesis possible.
Dr. E. T. Papoutsakis, for support, guidance and encouragement during the course of this research.
Dr. L. V. Inlntire and Dr. R. Storck, for serving on my thesis committee.
Chris Bussineau, for his assistance in the early part of this research.
Anil Diwan, for building the apparatus and develop¬ ing the procedure for the substrate-transport experiments, and for his generous help during the course of my work.
Monsanto Corporation, for providing me with the opportunity and financial support to make this endeavor possible. TABLE OF CONTENTS
Page
LIST OF TABLES vi
LIST OF FIGURES viii
NOMENCLATURES ix
ABSTRACT iii
INTRODUCTION 1
MATERIALS AND METHODS 14
I. Chemicals and Biochemicals 14 II. Chemical Assays 15 III. Selection and Growth of the Microorgan¬ ism 16 IV. Dry Weight Determination 20 V. Preparation of Cell-Free Extracts ... 21 VI. Enzyme Assays 21 VII. Yield Measurements 23 VIII. Other Characterization Techniques ... 24 IX. Preparation of Whole Cell Suspensions . 26 X. Transport Studies 26
RESULTS AND DISCUSSION 32
I. Isolation and Selection of the Micro¬ organism .... 32 II. Other Enzyme Activities for T15 .... 35 i) 3-Hexulose Phosphate Synthase (HPS ) 36 ii) Methanol Dehydrognase (MDH) ... 37 iii) Glucose 6-phosphate Dehydrogenase (GPD) and 6-Phosphogluconate Dehydrogenase (PGD) ...... 38
III. T15 Batch Growth Experiments 40 i) pH Profile 40 ii) Growth on Pure Substrates .... 42 iii) Growth on Mixed Substrates. ... 46 V
Page
iv) Discussion: The Mechanism of Substrate Inhibition 48 v) T15 Batch Yields 52
IV. Systematics: General Characterization of T15 58 i) General Characterization 59 ii) Morphological Characterization . . 59 iii) Physiological Tests. 60
V. Continuous Culture Experiments with T15. 62
VI. Formate Uptake by T15: Proton Transloca¬ tion Measurements 68
VII. Radiolabelled Substrate-Transport Studies 76 i) Measurement of T15 Cell Volume . . 76 ii) ^C-labelled Methanol Uptake ... 80 iii) 1’C-labelled Formaldehye Uptake. . 80 iv) l^C-labelled Formate Uptake. ... 84 v) Discussion 86
CONCLUSIONS AND RECOMMENDATIONS FOR FUTURE WORK . 93
LIST OF REFERENCES 97
APPENDIX 102 LIST OF TABLES
Table Page
1. NAD+-linked Formaldehyde and Formate Dehydrogenase Activities of Soil Isolates 34
2. T15 Growth on Various Carbon Sources 60
Appendix Table
A1. Formaldehyde DH Activities for Isolated Strains 102
A2. Formate DH Activities for Isolated Strains 103
A3. T15 3-Hexulose Phosphate Synthase Activities 104
A4. T15 MeOH Dehydrogenase Activities 106
A5. T15 Glucose-6-Phosphate Dehydrogenase Activities 107
A6. T15 6-Phosphogluconate Dehydrogenase Activities 108
A7. T15 Batch Yield Data 110
A8. Continuous Culture Yields for T15 Ill
A9. Glycerol Uptake Experiment #1 117
A10. Glycerol Uptake Experiment #2 118
All. Sucrose Uptake Experiment #1 119
A12. Sucrose Uptake Experiment #2 120
A13. Methanol Uptake Experiment #1 121
A14. Methanol UPtake Experiment #2 122
A15. Methanol Uptake Experiment #3 123 Table Page
A16. Formaldehyde Uptake Experiment #1 124
A17. Formaldehyde Uptake Experiment #2 125
A18. Formaldehyde + KSCN Uptake Experiment .... 126
A19. Formaldehyde + FCCP Experiment 127
A20. Formate Uptake Experiment 128
A21. Formate + KSCN Experiment 129
A22. Formate + FCCP Experiment #1 130
A23. Formate + FCCP Experimetn #2 131 LIST OF FIGURES
Figure Page
1. Oxidation Pathways in the Ribulose Monophosphate Cycle 4
2. The T15 pH profile 41
3. Batch growth curves for T15 at different initial methanol concentrations 43
4. The effect of fast transfers on T15 growth. . 45
5. The inhibitory effect of formaldehyde on T15 growth 47
6. Batch growth curves with methanol and formate 49
7. Biomass yield as a function of initial methanol concentration from batch experi¬ ments 53
8. Formate-induced proton uptake by T15 cells as a function of formate concentration. 72
9. The pH dependence of formate-induced proton uptake 74
10. Glycerol uptake profile 78
11. Sucrose uptake profile 85
12. Methanol uptake profile 81
13. Formaldehyde uptake profile 83
14. Formate uptake profile 85
Appendix Figure Page
Al. HPS activity as a function of reaction time . 105
A2. T15 cell dry weight determination 109 NOMENCLATURE
ADP,ATP Adenosine di-triphosphate BTB Bromothymol blue indicator CCCP Carbonyl cyanide m-chlorophenyl hydrazone D Dilution rate DCPIP 2,6-dichlorophenol indophenol
ddH20 Deionized, distilled water FCCP Carbonyl cyanide p-(triflouromethoxy)- phenylhydrazone FDDH Formaldehyde dehydrogenase FDH Formate dehydrogenase G PD Glucose-6-phosphate dehydrogenase HPS 3-hexulose phosphate synthase KSCN Thiocyanate MDH Methanol dehydrogenase
NAD,NADH2 Nicotinamide adenine dinucleotide and its reduced form O.D. Optical density PGA 3-phosphoglycerate PGD 6-phosphogluconate dehydrogenase PGI Phosphoglucoisomerase PHI Phospho-3-hexulose isomerase pmf Proton-motive force PMS Phenazine methosulfate P/O Moles of inorganic phosphate recovered per atom of oxygen taken up RMP Ribulose monophosphate S Substrate concentration SCP Single cell protein X Biomass concentration X P/0 ratio for methanol oxidation X
P/0 ratio for formaldehyde oxidation to formate Cell mass yield, g cells/ g substrate P/0 ratio for formate oxidation Specific growth rate 1
INTRODUCTION
In the last twenty years, a vast amount of litera¬ ture has been generated on the physiology and biochemistry of methylotrophic bacteria. Methylotrophs are defined as microorganisms which are able to grow at the expense of reduced carbon compounds containing one or more carbon atoms but no carbon-carbon bonds (Colby and Zatman, 1972).
Of special interest are those methylotrophic bacteria which grow on reduced one-carbon (C^) compounds, as they are ubiquitous in nature and are readily isolated from almost any sample of soil or water. Also, these bacteria play a vital role in the global biological carbon and nitrogen cycles. Aside from these fundamental character¬ istics, it was the appreciation of the potential of these bacteria for the production of single cell protein
(SCP) from methanol which stimulated the isolation and study of many new species (Anthony, 1982). Methanol is superior to other substrates for SCP prodution due in part to the attractive industrial characteristics that it possesses. MeOH is inexpensive, is produced commercially in large amounts from a variety of chemical feedstocks, is very pure, and is completely miscible in water
(Papoutsakis, 1976). Therefore, the methanol-utilizing 2 methylotrophic species are the most attractive for com¬ mercial exploitation.
More than half of the operating cost in any SCP process is that of the carbon substrate (Cooney, 1975); consequently, maximization of the biomass yield (Yx/S) is the primary objective of any biochemical engineer working with this process. Perhaps the single most important factor which influences the biomass yield of a methylotrophic species is the biochemical pathway through which the substrate carbon is incorporated into biomass. Carbon assimilation occurs by means of three primary reaction schemes in methanol-utilizing methylo- trophs? the ribulose biphosphate, serine, and ribulose monophosphate pathways. Anthony (1982) has provided a detailed summary and description of the biochemical reactions in each of these pathways. In the ribulose biphosphate pathway, or Calvin cycle, all of the carbon is assimilated at the oxidation level of CO2.
The serine pathway involves the formation of cell material from both formaldehyde and CO2. In the ribulose monophos¬ phate pathway (RMP) all of the cell carbon is assimilated at the oxidation level of formaldehyde. As a result, the RMP is the most efficient pathway since formaldehyde is at the same oxidation level as the cell material, and no reducing power is necessary for biosynthesis (Brock, 3
1979). Although ATP is required for carbon assimilation in all three of the pathways, this requirement is much lower with the RMP pathway. These bioenergetic advan¬ tages result in higher biomass yields for RMP bacteria; consequently, methanol-utilizing RMP -species are the microorganisms of choice for SCP production.
Methylotrophs derive most of their biosynthetic energy through the oxidation of their substrates. In RMP bacteria, the complete oxidation of methanol to CO2 can be accomplished through two different routes, which are diagrammed in Figure 1 (Strom et al., 1974). In the linear route, the oxidation is catalyzed from methanol to formaldehyde to formate to CO2 by three dehydrogenase enzymes: methanol dehydrogenase (MDH), formaldehyde dehydrogenase (FDDH) and formate dehydrogenase (FDH).
These enzymes are not unique to RMP bacteria? in fact, many methylotrophs are able to oxidize methanol by this simple linear scheme. The cyclic oxidation route shown in Figure 1, on the other hand, is Unique to RMP species.
In the first reaction, 3-hexulose phosphate synthase
(HPS) catalyzes the aldol condensation of formaldehyde and ribulose-5-phosphate to give hexulose-6-phosphate, which subsequently undergoes two isomerization reactions to form glucose-6-phosphate. Glucose-6-phosphate dehydrogenase (GPD), an NAD-linked enzyme, catalyzes the MDH FDDH CH,OH > CHzO > HCOOH ——> C02
Figure 1. Oxidation pathways in the Ribulose Monophosphate Cycle 4 5 oxidation of glucose-6-phosphate to 6-phosphogluconate.
As Figure 1 shows, it is through this key intermediate that assimilation of carbon into biomass occurs.
Therefore, several of the enzymes in the cyclic oxidation route, namely HPS, PHI, PGI and GPD, are also part of the normal assimilatory pathway of RMP-cycle bacteria. The most important enzyme of the cyclic oxidation route is 6- phosphogluconate dehydrogenase (PGD), which catalyzes the irreversible oxidative decarboxylation of 6-phospho¬ gluconate to ribulose-5-phosphate, thus completing the cycle. PGD activity, therefore, results in the branching of the carbon flow from assimilation to oxidation, and for this reason the cyclic oxidation route is sometimes called the dissimilatory cycle of formaldehyde oxidation
(Anthony, 1982). The cyclic and linear oxidation routes operate simultaneously in RMP bacteria in unknown extents each. Some researchers have suggested that formaldehyde
is oxidized primarily by the dissimilatory pathway
(Anthony, 1982; Zatman, 1981), while others have provided experimental evidence that the linear route is the primary mechanism of substrate oxidation (Ben-Bassat et al.,
1980, Papoutsakis and Lim, 1981). The branching of the carbon flow from one oxidation route to the other is most
likely influenced by a number of environmental, physiological and biochemical factors. 6
The reactions of the linear oxidation route are of primary importance due to their potential for ATP production. Methanol oxidation to formaldehyde is catalyzed by an NAD-independent alcohol dehydrogenase, and the natural electron acceptor of this reaction remains unknown. It has been suggested that the enzyme is a membrane-bound pteridoprotein that interacts with the electron transport system directly at the level of cytochrome c (Anthony, 1978). Direct experimental measurements of the P/0 ratio for MeOH oxidation by a number of researchers (Tonge et al., 1977; van Verseveld and Stouthamer, 1978; Drozd and Wren, 1980) have indicated that only one mole of ATP is formed per mole of methanol oxidized; therefore, methylotropic bacteria exhibit a very low efficiency (~20%) of energy conservation at this step in terms of the available energy of the reaction.
The standard redox potential of the CH3OH/HCHO half reaction (-.182 V at pH 7.0) is low enough to couple at the flavoprotein level, which would give a P/O ratio of
2. Although this value is greater than the experimentally observed value of P/O =1, it is interesting at least to note that 2 moles of ATP could potentially be produced from the oxidation of 1 mole of MeOH. This value, therefore, represents an upper limit for methanol oxida¬ tion • The production of 3 ATPs from the oxidation of 7
MeOH (corresponding to a P/0 ratio of 3) is highly unlikely because the redox potential is not sufficiently negative for the reduction of NAD (Ribbons et al., 1970).
An NAD-linked formaldehyde dehydrogenase operates in all serine pathway bacteria and in some RMP bacteria. Several
investigators have also found NAD-independent FDDH activity in RMP species (Johnson and Ouayle, 1964; Hirt et al.,
1978; Patel et al., 1980b). Formate dehydrogenase, on the other hand, is always NAD-linked. In the case of the NAD-dependent enzymes, electrons are passed from the substrate to NAD+, and the subsequent oxidation of
NADH2 via the electron transport chain facilitates the generation of ATP by thé membrane-bound ATPase. Published experimental evidence suggests maximum P/O ratios of 3 for the formaldehyde and formate oxidation steps (Hammond and Higgins, 1978; van Verseveld and Stouthamer, 1978).
Therefore, three moles of ATP can apparently be formed
from the oxidation of one mole of formaldehyde or formate
(NAD-linked only) and the efficiency of energy coupling
for these two oxidation reactions is much higher than
that for the methanol oxidation. This experimental
evidence suggests that FDDH and FDH are key enzymes for
the production of biosynthetic energy.
In addition to direct experimental measurements of
P/O ratios, a number of researchers have computed biomass 8 yields based upon theoretical predictions and assumptions
involving the cell ATP yield (van Dijken and Harder,
1975; Anthony, 1978). These theoretical developments typically require the estimation of P/0 ratios for substrate oxidation; therefore, a comparison of computed yields with experimental yields for a particular species provides an indication of the actual ATP yield from each oxidation step. For example, in his studies with
Methylomonas EP-1, a RMP-cycle methanol-utilizing bact¬ erium, Papoutsakis (1978a) has reported very high experimental biomass yields of up to 0.65 g cells/g MeOH.
In a theoretical computation of the yield for this species (Papoutsakis and Lim, 1981), the assumption of P/O ratios of 1, 3 and 3 for MeOH, formaldehyde and
formate oxidation, respectively, best suited the experi¬ mental findings (computed yield = 0.63). Theoretical developments such as this provide additional evidence
that FDDH and FDH are of particular importance for ATP
production. In theory, a species with high activities
of these two enzymes would be capable of producing more
ATP; consequently, more energy would be available for
assimilation of carbon into biomass. It seems reason¬
able that such a species would be bioenergetically superior
and would give higher biomass yields than species
with lower FDDH and FHD activities. Indeed, the EP-1
strain mentioned earlier displayed relatively high fn 9 vitro activities for both of these enzymes, and the reported biomass yields of ~0.65 were higher than any other reported in the literature (Papoutsakis, 1976).
The principles and results described above provided the basis for the research in this thesis. Several
MeOH-utilizing strains were isolated from soil, and enzyme assays were performed in order to determine the in vitro FDDH and FDH activities. Eventually a strain, designated T15, was selected for further study on the basis of high activities for these two key enzymes. In order to test the validity of the selection criteria, the growth of the new strain was studied in both batch and continuous culture. In addition, a variety of biochemical and microbiological techniques were used to further characterize the strain. The detailed results of these studies are presented in subsequent sections of this thesis.
Another primary focus of this research has involved the study of the substrate-transport mechanisms of the
T15 strain. Substrate-transport in bacteria is accom¬ plished through the following four mechanisms (Wang et al., 1979):
1. Passive Diffusion: Entry is by random molecular
motion. This process does not require energy
and does not lead to concentration against a
gradient. 10
2. Facilitated Diffusion: The solute combines
reversibly with a specific carrier protein in
the membrane which oscillates between the inner
and outer surface. The carrier binds the solute
on the outside and releases it on the inside
of the cell. This process does not require
energy or lead to concentration against a
gradient.
3. Group Translocation: The solute is covalently
modified, usually by phosphorylation, and this
reaction facilitates passage of the modified
solute across the membrane. This process
requires energy.
4. Active Transport: This process involves a
complex membrane transport system which is
highly specific for the solute. This process
is coupled to the expenditure of metabolic energy.
Accumulation of a solute against a concentration
gradient is indicative of active transport.
Active transport is perhaps the most important mechanism due to its energy requirement.
The chemiosmotic hypothesis of Peter Mitchell serves as the basis for a complete understanding of the bioenergetics of active transport. In his early studies of mitochondrial membrane systems, Mitchell showed that the 11 primary event resulting from the transfer of electrons down the membrane-bound respiratory chain was the translocation of protons and hydroxyl ions to opposite sides of the membrane (Mitchell and Moyle, 1967). Such a separation results in a combined chemical gradient of hydrogen ions (pH gradient) and electrical potential gradient across the membrane. This electrochemical gradient, sometimes called the proton-motive force (pmf or
Ap may be expressed mathematically as follows (Rosen and
Kashket, 1979):
p = A* - ZApH (1) where Z = 2.3RT/F, F = Faraday constant
A'fc = electrical gradient
ApH = chemical gradient of protons
In bacteria, the cytoplasmic membrane contains the electron transport chain, and respiration results in the flow of protons to the outside of the membrane. In addition to respiration-linked proton transport, a proton-motive force can also be generated by means of
ATP hydrolysis. Once the pmf is generated, it is used by bacteria to transport nutrients across the cytoplasmic membrane. The work of active transport is done at the level of the membrane-bound carrier proteins, also called 12 permeases. Either component of the proton-motive force
(A'I'or ApH) can drive active transport; alternatively, the total pmf may serve as the driving force.
Very little is known about the bacterial transport systems for one-carbon compounds. Typically it is assumed that small, neutral molecules such as methanol and formaldehyde are transported across the cell membrane by passive diffusion (Anthony, 1982; Bellion et al.,
1983). These molecules have a high permeability (PcH30H =
2.26 x 10~^cm/s; Stein, 1967), so the assumption of diffusion at first seems reasonable; however, preliminary experimental observations with methylotrophic bacteria have indicated the existence of asymmetric transport
systems. For example, experiments with Methylomonas L3
cells have shown internal concentrations of formaldehyde
from 10 to 90 times that of the external concentration
(Krug et al., 1979). Also, in order to maintain the
fast growth rates observed in continuous culture with methylotrophic bacteria (washout dilution rates in excess
of 0.5 hr“l; Papoutsakis, 1976; Hirt et al., 1978),
the accumulation of methanol and formaldehyde is apparently
necessary, since the external concentrations of these
intermediates at steady-state are approximately zero.
In addition, the high iji vitro activities reported for
the key oxidative enzymes (Papoutsakis, 1976; Krug et 13 al., 1979; Beinor, 1978) suggest that these bacteria maintain high internal levels of the substrates.
The presence of active transport systems as sug¬ gested by this preliminary evidence was finally confirmed by Diwan et al. (1983). In his work with Methylomonas
L3, Diwan identified active transport systems for both methanol and formaldehyde. Using radiolabelled sub¬ strates, Diwan was able to measure up to a 10-fold accumulation of these substrates in the cells. These asymmetric transport systems were found to have a strong effect on the transient behavior of the chemostatic cultures. These results provided the basis for the substrate-transport experiments with the T15 strain as described in this thesis. 14
MATERIALS AND METHODS
I. Chemicals and Biochemicals
The growth medium for the bacteria was composed of a mixture of analytical grade salts purchased from com¬ mercial sources. Bacto-Agar from Difco Laboratories and
Standard Methods Agar from Becton Dickinson Co. were used in the preparation of the agar plates and slopes. The formaldehyde and formate solutions were prepared from practical-grade paraformaldehyde (Papoutsakis, 1976) and potassium formate purchased from the Eastman Kodak
Company. Methanol solutions were prepared by dilution of 100.0% Absolute MeOH from Baker Chemicals with deionized, distilled water.
The phosphate, tris-HCl, KC1, KCl-glycylglycine, and triethanolamine buffer solutions were prepared by diluting appropriate quantities of the buffer salt with deionized, distilled water and subsequently adjusting the pH with concentrated HC1 or NaOH solutions. All buffer salts were purchased from commercial sources.
The chromotropic acid, 2-mercaptoethanol, human serum albumin, carbohydrates, amino acids, enzymes, coenzymes, ionophores and drugs used in the various assays and transport studies were all obtained from the Sigma 15
Chemical Company. The l^C-labelled methanol, formate and
formaldehyde used in the substrate-transport experiments were purchased from either Amersham or ICN.
II. Chemical Assays
Methanol concentrations were assayed by means of gas
chromatography with an Antek Series 300 GC equipped with
a flame ionization detector. The 2-1/2 ft long, 1/8 inch
I.D. column was packed with Porapak 0, 50-80 mesh.
Operating temperatures were 200°C for the injection port
and detector, and 140°C for the column. The hydrogen and
air (detector gases) flow rates were 25 cc/min and 250
cc/min, respectively, while the helium (carrier gas) flow
rate was regulated at 30 cc/min. The sample injection
volume remained constant at 1.0 ul. The detector output
signals were recorded and integrated with a Cole Palmer
Model 8384-32 chart recorder.
Formaldehyde concentrations were determined by means
of the colorimetric assay described by West and Sen
(1956). First, a 1% (w/v) mixture of chromotrophic acid
in concentrated reagent grade H2SO4 was prepared. Next,
1 ml assay samples, together with formaldehyde standards,
were combined with 1 ml aliquots of the chromotropic
acid mixture and another 10 ml of concentrated H2SO4.
The resulting mixtures were quickly vortexed, and after
30 min. their extinction was measured at 570 nm with a 16
Bausch and Lomb Spectronic 20 spectrophotometer. This method was accurate in the range from 0.2 to 20.0 ug/ml of formaldehyde.
Protein concentrations were assayed by the Folin-
Ciocalteau method, also known as the Lowry protein assay
(Lowry et al., 1951). Human serum albumin was used as the protein standard.
Ill. Selection and Growth of the Microorganism
The original objective of this work was to select a methylotrophic strain with similar enzymatic activities to Methylomonas EP-1, as mentioned earlier; consequently, the medium used for the selection and growth of T15 was identical to that used in the work with EP-1 (Papoutsakis,
1976). This medium was a modified version of the basal
L-salts medium described by Whittenbury, et al. (1970).
The final composition was as follows; MgS04*7H20 1.0
7 g/1, FeS04’ H20 0.01 g/1, CaCl2 0.1 g/1, NH4C1 0.3 g/1, ZnS04*7H20 0.025 g/1, EDTA-disodium salt 0.004 g/1, MnS04*H20 10 ug/1, Na2Mo04*2 H20 10 ug/1,
CUS04*5H20 5ug/l, COC12*6H20 10 ug/1, H3P04 10 ug/1, and 60 ml/1 of pH 6.9, 0.6 M phosphate buffer.
Deionized, distilled water was used in the preparation of all media. The salts medium was steam sterilized in an autoclave for all batch experiments, while in continuous 17 culture experiments, which required large volumes of media, the salts solution was filter sterilized twice
through 0.2 u Gelman filters. For agar plates, 15.9 g/1 of solid agar was mixed with the heated salts solution prior to the sterilization. The substrates (methanol,
formaldehyde, formate, etc.) were added aseptically to
the sterilized medium in the desired concentrations.
The first step in the selection of the T15 strain
involved the treatment of organic compost, a typical
environment for methylotrophic bacteria (Anthony, 1982),
with a dilute methanol solution. The methanol was
sprinkled onto the compost periodically for about one
month, after which a sample of the treated soil was placed
into a 2L Erlenmeyer flask containing 500 ml of the
sterilized liquid salts medium and 0.2% (v/v) MeOH.
This culture was then incubated at 30°C in a New Brunswick
Controlled Environment Incubator Shaker at a shaker speed
of 270 rpm. After a period of about 24 hrs the liquid
became turbid with bacterial growth, and agar plates
containing 2% (v/v) MeOH were streaked with the suspension.
The plates were incubated for several days at 30°C in a
controlled environmental chamber, and a microscopic
examination revealed several colonies which differed
significantly in size, color and other morphological
characteristics. Several pure cultures were isolated 18 from these mixed cultures, and eventually T15 was selected as the strain for further study on the basis of key enzyme activities. A more detailed account of the isolation and selection procedure is given in the Results and Discussion section.
All subsequent batch culture experiments with T15 were performed in 250 or 2000 ml flasks, with working volumes of 50 and 500 ml, respectively. The incubator shaker conditions described earlier remained constant in all of these studies.
Continuous culture experiments were conducted with a
New Brunswick Model C-30 Bioflo Chemostat. The 1L vessel had a working volume of 350 ml and contained an overflow tube for level control. The medium flow (dilution rate) was contolled by means of a medium feed pump built into the unit. The pH of the culture was monitored with an
Ingold pH electrode and was controlled at pH 7.0 with a
Horizon pH controller. A IN NaOH solution was used for pH control. Dissolved oxygen levels were measured with a galvanic D.O. probe and recorded by a New Brunswick D.O. monitor. The agitation rate remained constant at 500 rpm, and the temperature was controlled at 30°C by means of an immersible heater. The reactor was inoculated from batch liquid cultures, and it was operated in a batchwise mode until the cell suspension in the reactor reached a 19 turbidity of 0.5 to 0.6 O.D. at 600 nm. At this point, continuous operation was initiated.
The bacterial growth in all liquid cultures was monitored with a Gilford Model 250 Spectrophotometer at either 550 or 600 nm. Appropriate blanks were used to correct for the precipitation of phosphates in the suspension. Alternatively, dilutions were made with a low pH saline solution in order to eliminate any precipi¬ tate interference.
Long term preservation of T15 was accomplished by lyophylization. Cells were grown in a 500 ml batch culture on 0.2% (v/v) MeOH until late exponential phase.
The cells were then harvested by centrifugation for 10 min at 12,000 rpm, and the pellet was resuspended in 10 ml of a 25% (v/v) skim milk/dd H2O solution (cryoprotective agent) which had previously been steam sterilized. A 2 ml aliquot of the final suspension was placed into each freeze-drying vial, and the cells were quick-frozen into a shell in a dry ice/acetone bath. The vials were then placed in the lyophylizer under a vacuum. When the cells were completely dry, the vials were removed and vacuum sealed, and the cultures were stored in the laboratory freezer. Although the rehydrated cells exhibited a prolonged lag phase, upon extended incubation the major¬
ity of the lyophylized cultures proved to be viable. 20
IV. Dry Weight Determination
Cells grown in batch culture with 0.2% (v/v) MeOH were harvested, and the pH of the suspension was lowered to 5.0 with concentrated HC1 in order to dissolve the phosphates. The suspension was then centrifuged at
11,000 rpm in a Sorval RC-5B Refrigerated Centrifuge (GSA rotor), and the cells were subsequently washed with a
1.5 g NaCl/L pH 5.0 saline solution. This centrifugation procedure was repeated twice, and the final pellet was resuspended in a 0.5 g NaCl/L, pH 5.0 saline solution.
Dilutions of the final suspension were prepared in tripli¬ cate, and their optical densities were measured at 600 nm with the Gilford-250 Spectrophotometer. Seven ml aliquots of the final homogeneous suspension were then placed into four 10ml pre-weighed Sorval glass centrifuge tubes, and these were centrifuged at 15,000 rpm (SS-34 rotor) for 30 min. All of the above operations were carried out at 0 to 4°C. The supernatant was then dis¬ carded, and the cell pellets were dried in an oven at
104°C for 48 hrs (or until a stable weight was obtained).
The pellets were then dessicated over phosphorous pentoxide for 24 hrs., and afterwards the tubes containing the dried pellets were reweighed. The accuracy of this method was better than 0.4% for the quatriplicate samples. 21
V. Preparation of Cell-Free Extracts
Cells grown in batch culture on 0.1% (v/v) MeOH were harvested by centrifugation at 7500 rpm (55-34 rotor) for
20 min. The pellets were resuspended in 0.2M, pH 7.0 phosphate buffer plus 2mM 2-mercaptoethanol. The centrifugation and washing procedures were repeated twice in order to remove any residual substrate. The final pellet was resuspended in 3-5 ml of buffer per gram wet weight of cells. The suspension was then placed into aluminum cooling cells, and the cell membranes were dis¬ rupted by sonication for approximately 4 min. using a
Heat Systems Ultrasonics Model W-140 Sonifier. The disrupted cells were then centrifuged at 18,000 rpm in order to remove the cell debris. The resulting super¬ natant was used as the cell-free extract in all of the enzyme assays. All of the above operations were per¬ formed at 0-4°C.
VI. Enzyme Assays
A Gilford Model 250 Spectrophotometer and Model 6051
Thermal Chart Recorder were used for all spectrophoto- metric assays. The rates of substrate utilization generally decreased with time; consequently, only initial rates were used for activity calculations.
i) Methanol Dehydrogenase
PMS and DCPIP linked activity was assayed at pH 9.5 22
according to the method of Anthony and Zatman (1967).
Reagent concentrations were as follows: 300 umoles Tris-
HC1 buffer, pH 9.5, 45 umoles NH4CI, 3 umoles KCN, 0.13
umoles DCPIP, 0.33 umoles PMS and 16 umoles MeOH. Cell
extract and ddH20 were added to the reaction mixture to
bring the total assay volume to 3 ml. The reduction of
the artificial electron acceptor in the assay mixture
resulted in a decrease in visible absorbance at 600 nm.
ii) Formaldehyde Dehydrogenase and Formate
Dehydrogenase
NAD-linked activity was assayed with a reaction
system similar to that described by Johnson and Quayle
(1964). The assay mixture contained 1.5 umoles NAD, 300
umoles 0.6 M pH 7.5 phosphate buffer, 15 umoles formal¬
dehyde or 42 umoles formate, and 0.4 or 0.6 ml cell
extract. The formaldehyde dehydrogenase system also
contained 4 umoles GSH. Reduction of the NAD was
monitored as an increase in UV absorbance at 340 nm.
iii) 3-Hexulose Phosphate Synthase
The procedure was a modified version of the method
described by Lawrence et al. (1970). The reaction
mixture contained 300 umoles 0.6M, pH 7.0 phosphate
buffer, 25 umoles MgCl2» 25 umoles D-ribose-5-phosphate,
25 umoles Formaldehyde, 1.2 ml H2O and 0.3 ml cell-free
extract 23
The total assay volume was 5 ml. The reaction was started upon addition of the cell extract to the mixture and was stopped through the addition of 0.3 ml of 4.0M
HC1. A 5ml aliquot of 20% TCA was subsequently added in order to precipitate the protein, and the mixture was centrifuged at 15,000 rpm for 25 min. The supernatant was then assayed for the substrate uptake by means of the colorimetric formaldehyde assay as described earlier.
Total reaction times of 1,2,3,5 and 10 min. were studied.
iv) Glucose-6-Phosphate Dehydrogenase and 6-
Phosphoqluconate Dehydrogenase
NAD-linked activity was assayed using a modified system based on the reaction mixture developed by Hohorst
(1965). The assay mixture contained 4Ô0 umoles 0.4M, pH
7.6 Triethanolamine buffer, 50 umoles MgCl2, 1.5 umoles
NAD, 15 umoles 6-Phosphogluconate or 15 umoles Glucose-6-
Phosphate and 0.5 ml cell-free extract. An increase in
UV absorbance at 340 nm due to the reduction of NAD provided a measure of the enzyme activities.
VII. Yield Measurements
Cell yields (Yx/S, g cells/g MeOH) were measured
from batch cultures at MeOH concentrations ranging from
.025 to 3.25% (v/v). Samples were taken both before
inoculation as well as after the cultures had reached 24 stationary phase. The optical densities of the stationary phase samples were measured at 600 nm in order to determine the dry weight of cells. All samples were then filtered through 0.2u Gelman membrane filters in order to remove the cells. The filtrates were subsequently assayed for
MeOH content by means of gas chromatography as described earlier.
In continuous culture experiments the samples were taken at steady-state directly from the reactor, and the extinction of these suspensions were determined at 600 nm. Filtrates from reactor and feed tank samples were then assayed for MeOH.
VIII. Other Characterization Techniques
i) Gram Stain
Gram stain slides of the T15 bacteria were prepared
from batch liquid cultures using a Difco Gram Stain Kit.
Before staining, the cells were suspended in 5% (v/v)
formalin and concentrated by centrifugation in order to preserve the size and shape and to facilitate solid
staining (Murray, et. al. 1981).
ii) Catalase Test
A few drops of 3% hydrogen peroxide were applied to
the bacterial colonies on agar plates. These colonies
were then examined microscopically for the evolution of 25 bubblies, which indicated a positive response (Smibert and
Krieg, 1981). iii) Oxidase Test
A piece of filter paper was moistened with a few drops of 1% tetramethyl-p-phenylenediamine dihydrochloride, and growth from the surface of an agar plate was streaked onto the moistened paper. A positive test was indicated by the development of a violet or purple color in 10 sec
(Smibert and Krieg, 1981). iv) Growth and Acid Production on Various Carbon Sources
Media for testing other carbon sources contained the basal salts in the concentrations stated previously, 0.1%
(v/v) of bromothymol blue (BTB) indicator, and 0.1% (w/v) of each carbon source (Chen et al., 1977). The carbon sources were prepared as concentrated solutions and filter sterilized, while the salts medium containing the
BTB indicator was placed into standard fermentation tubes in 10 ml aliquots and autoclaved. The tubes were inoculated with 0.1 ml of late exponential phase T15 bacteria. The production of acid was indicated by a change in the color of the BTB dye from green to yellow.
Growth on the various carbon sources was indicated by an increase of turbidity in the fermentation tubes. Tubes containing MeOH as the sole carbon source were used as a 26
control. Nutrient agar plates were also inoculated with
T15 in order to examine the growth on complex media.
IX. Preparation of Whole Cell Suspensions
Whole cell suspensions of T15 for transport studies were prepared according to the method described by West
and Mitchell (1973). A small inoculum from an agar plate was grown for 48 hrs at 30°C in a 50 ml batch liquid
culture (0.1% v/v MeOH). Small samples from this culture were then used to inoculate two 500 ml batch cultures with 0.1% v/v MeOH. The stationary phase of growth,
corresponding to an OD at 600 nm of about 0.9, was
reached one hour before harvesting. The cells were
harvested by centrifugation at 4°C for 10 min at 12000
rpm, washed twice with a pH 7.0 140 mM KC1 solution
buffered with 1.5 mM glycylglycine, and resuspended at a
cell density of 30-50 mg dry wt./ml in the same solution.
Cells were resuspened in unbuffered 140 mM KC1 if sensi¬
tive pH measurements were required.
X. Transport Studies
i) Experimental Apparatus
The apparatus used for the transport studies with
T15 was developed by Anil Diwan in his studies with
Methylomonas L3 (Diwan, et al., 1983). A Yellow Springs
Instruments Co. (Ohio) Model 53 bath assembly system was 27 used for the incubation of the T15 cells. The cell suspension was placed into four 15 ml glass sample chambers which were submerged in a cylindrical glass vessel. A
VWR Scientific constant temperature circulating water bath was used to maintain a constant temperature of 33°C
in the vessel. The sample chambers contained stirring discs which were driven by a magnetic stirring unit built
into the YSI system. The stirrer speed remained constant at 400 rpm in all experiments. Lucite plungers, which provided a barrier to oxygen diffusion, were placed in the sample chambers above the cell suspensions. These plungers were machined to accommodate oxygen and pH probes. Each plunger also contained an access slot which
facilitated the injection of chemicals into and removal of samples from the stirred suspension. SMI micro¬ pipettors were used for the transfer of materials to and
from the sample chambers.
The YSI system also included a biological oxygen monitor with two polarographic oxygen electrodes. Each
electrode consisted of a platinum cathode and a silver
anode encased in carbon black. A thin teflon membrane
isolated the sensor elements from the environment. This 28 system was capable of measuring oxygen consumption rates as low at 7 ul 02/hr.
A Radiometer Copenhagen PHM 64 research pH meter was used for sensitive pH measurements in the T15 suspensions.
The system included two RC combined glass pH electrodes which were capable of measuring pH changes within + 0.002 pH units.
Output signals from the oxygen and pH meters were simultaneously recorded with a Hewlett Packard Model 7100BM strip chart recorder. This instrument was equipped with input range and attenuation controls in order to adjust the amplitude and offset of the input signals.
ii) Proton Translocation Measurements
The uptake of protons accompanying the transport of formate by T15 cells was studied in the experimental apparatus described above. Freshly prepared whole cell suspensions of T15 in 140 mM KC1 (unbuffered solution) were incubated in the New Brunswick shaker for 1 hr in order to consume as much of the endogenous substrate as possible. During this time, the pH and O2 electrodes were calibrated, and the water bath was started in order to allow temperature equilibration to 33°C. Approximately
10 ml of cell suspension was then placed into each sample chamber, and the plungers, O2 electrodes and pH electrodes were put into place. After constant pH and O2 slopes were 29 obtained (10-15 min), the pH scale on the chart recorder was calibrated by injecting 10 ul of 5N HC1 and monitor¬ ing the pen deflection. The pH of the suspension was adjusted to the desired value with 100 mN HC1 or 100 mN
NaOH. Solutions of sodium or potassium formate, in concentrations ranging from 25 mM and 1M, were then injected into the sample chambers through the plunger access slots. The changes in pH and oxygen concentration resulting from the uptake and oxidation of the formate were monitored on the chart recorder.
iii) Radiolabelled Substrate Uptake Experiments
The method used for the labelled substrate uptake experiments with T15 was identical to that used by Diwan et al. (1983). Six ml aliquots of the T15 whole cell suspension in KCl/glyclyglycine were placed in the YSI sample chambers, and the cells were aerated for 1.5 hr in order to consume the endogenous substrate. The plungers,
C>2 electrodes and pH electrodes were then put into place, and the cells were allowed to go anaerobic (2-3 hrs). The access slots in the plungers were flushed with nitrogen in order to reduce the leakage or air into the suspen¬ sions. Once the cells had become anaerobic, the pH of the suspension was adjusted to pH 7.0, and after 5 min had passed the l^C-labelled methanol, formaldehyde or formate was injected into the sample chamber in the 30 desired concentration. Ten 50 ul samples were then quickly withdrawn and placed into 25 mm Nucleopore Swin- lok filter holders containing 0.4u pre-wetted poly¬ carbonate filters. After each sample was taken, 5 cc of air (in a syringe) was immediately passed through the filter in order to push out the adhering liquid. The filter holders had previously been placed onto scintilla¬ tion vials in order to collect the supernatant. Two additional unfiltered samples were taken in order to measure the total activity. After all of the samples had been flushed with air, 3 cc of the KCl/glycylglycine solution was passed through the filters in order to wash the adhering radioactivity from the cells. The wash solution was kept at room temperature to avoid the loss of internal substrate due to temperature transition
(Leder, 1972). Another 5cc of air was then immediately flushed through the filters in order to force the wash liquid into the scintillation vials. 100 ul aliquots of
2N KOH were placed onto the filters and into the super¬ natant vials in order to kill the cells and avoid any further metabolism. The filters were then removed from the holders and placed into ten new scintillation vials.
ACS xylene-based scintillation fluid was added to each of the vials, and the samples were counted in a Beckman liquid scintillation counter. The polycarbonate filters 31 were soluble in the scintillant so that the filter
samples, as well as the supernatant samples, were
completely homogeneous.
In some experiments, FCCP or KSCN was incubated with
the cell suspension for 10 min prior to the injection of
the labelled substrate. 32
RESULTS AND DISCUSSION
I. Isolation and Selection of the Microorganism
In an effort to select for methanol-utilizing species, a sample of compost was treated with a dilute
(0.5% v/v) methanol solution over a period of several weeks. A sample of the soil was incubated in the liquid salts medium with 0.2% (v/v) MeOH until the growth became turbid. Agar plates containing 2% (v/v) MeOH were then streaked with this suspension, and a microscopic examina¬ tion of the growth on these plates revealed a mixed culture of several colonies with different morphological characteristics. Each of these different colonies was subcultured onto other agar plates by means of the common streak-plate method described by Krieg (1981) which
invariably yielded well isolated colonies. Through this process of serial subculturing and microscopic examination of the plates, three pure cultures were eventually isolated
(after about one month) from the original mixed culture.
The morphological characteristics of the colonies from
these three isolates, designated Til, T13 and T15, differed
significantly from each other, as the descriptions below
indicate. 33
Til Circular, convex colonies; Diameter ~1 nun. Colonies are smooth, white in color, and trans¬ lucent.
T13 Circular, convex colonies; Diameter ~1.2 mm. Colonies are smooth and light yellow in color. Characterized by a thick, yellow translucent center, surrounded by a clearer outside ring which is almost transparent.
T15 Circular, convex colonies; Diameter ~2 mm. Colonies are smooth, white in color, and trans¬ lucent. Characterized by the presence of bubbles in the colony which become larger near the center.
The descriptions above were made from plates of identical age (1 week) which had been inoculated from late exponen¬ tial phase liquid cultures of the three strains.
Incubation conditions for the plates were identical.
In addition to these morphological differences, preliminary batch growth experiments with the three isolated strains revealed different growth characteris¬ tics. The Til and T15 strains grew much faster in batch culture than the T13 strain. Also, T13 tended to form large aggregates, particularly in the early stages of batch growth.
As described in the Introduction, the NAD-linked formaldehyde and formate dehydrogenases of methylo- trophic bacteria are key enzymes for the production of biosynthetic energy. The FDDH and FDH enzyme assays were performed in an effort to select for a strain with high in vitro activities. Two separate batches of extract 34 were prepared from each strain, and the specific activi¬
ties listed in Table 1 represent average values. The
raw data and calculation procedure for the enzyme assays may be found in the Appendix.
Table 1. NAD+-linked Formaldehyde and Formate Dehydro¬ genase Activities of Soil Isolates
FDDH Specific Activity FDH Specific Activity nmoles nmoles Strain min-mg protein min-mg protein
Til 10.6 338.3
T13 31.3 17.6
T15 19.0 678.2
No dye-linked formaldehyde dehydrogenase activity (Johnson
and Quayle, 1964; Hirt et al., 1978) was found in any of
the extracts.
The specific activities of formaldehyde dehydrogenase
in crude extracts from different bacteria, as reported in
the literature (Johnson and Quayle, 1964; Colby and Zatman,
1973; Ferenci et al., 1975; Papoutsakis, 1976; Ben-Bassat
and Goldberg, 1977; Hirt et al., 1978; Stirling and
Dalton, 1978; Marison and Attwood, 1980; Anthony, 1982),
vary between 6 and 84.4 nmoles/min-mg protein, with the
majority of activities between 10 and 25 nmoles/min-mg 35 protein. These results reflect to some extent the various growth conditions, cell disruption methods, and methods of enzyme assay, but they also suggest an actual range of activities for methylotrophic bacteria. As Table 1 shows, the FDDH activities for the three isolates were in the expected range, with T13 showing a slightly higher activity than most methylotrophs.
Reported literature values for the specific activities of formate dehydrogenase range from 4 to 183 nmoles/min- mg protein (same references as for FDDH, plus Patel et al., 1978). The FDH activity for isolate T13 was in the expected range; however, the activities for Til and T15 were significantly higher than any of the reported literature values.
Based upon these enzyme activities, T15 was selected as the strain for further characterization and study.
The FDDH and FDH specific activities for T15 were twice
as large as those found for Til. T13 was not considered
to be a good candidate due to the relatively low activity
for FDH.
II. Other Enzyme Activities for T15
Several additional enzyme assays were performed with
T15 in order to characterize the carbon assimilation
pathway of the strain. 36
i) 3-Hexulose Phosphate Synthase (HPS)
This enzyme catalyzes the aldol condensation of formaldehyde with ribulose 5-phosphate to form 3-hexulose
6-phosphate in the first carbon fixation step of the ribulose monophosphate pathway. The presence or absence of this enzyme may be regarded as a positive or negative test for the involvement of the RMP cycle (Papoutsakis,
1976).
Reaction times from 1 to 10 min were studied. The resulting enzyme activity profile for T15 is shown in
Figure A1 in the Appendix. Typically, specific activities for HPS in the literature are reported as average values over the initial period of reaction time when the CH2O uptake is linear (Beinor, 1976). For T15, the reaction was linear for about the first 3 min. On this basis, the specific activity was as follows:
HPS Specific Activity = 263.9 umoles/hr*mg protein
Because of the position of HPS in the RMP carbon assimila¬ tion pathway (refer to Figure 1), the specific activity of HPS provides an indication of the efficiency of carbon incorporation into biomass (Papoutsakis, 1976). The HPS specific activity for T15 was among the highest of those reported in the literature (Lawrence et al., 1970; Ferenci et al., 1974; Sahm and Wagner, 1974; Strom et al., 1974; 37
Beinor, 1976; Papoutsakis, 1976; Sahm et al., 1976b,
Kato et al., 1978, Beardsmore et al., 1982). This result provided a preliminary indication that T15 was indeed an efficient strain.
ii) Methanol Dehydrogenase (MDH)
The oxidation of methanol to formaldehyde by methylotrophic bacteria is typically catalyzed by a NAD+- independent dehydrogenase, which was originally described by Anthony and Zatman (1964). The natural electron acceptor for this reaction remains unknown. There is substantial evidence, however, that MDH is a membrane- bound pteridoprotein which passes electrons directly to the electron transport chain at the level of cytochrome c
(Anthony, 1975b; Anthony, 1978b). In order to assay for the enzyme it is necessary to use the artificial electron acceptor phenazine methosulphate (PMS). The reoxidation of the reduced PMS in the assay is coupled to the reduction of the dye 2,6-dichlorophenol indophenol (DCPIP), which can be measured spectrophotometrically (Anthony and
Zatman, 1967).
The average specific activity in T15 extract was as
follows:
MDH Specific Activity = 8.56 nmoles/min-mg protein 38
Literature values for MDH in crude extracts range between
4 and 1300 nmol/min-mg .protein (Anthony, 1982). The observed activity for T15 lies at the low end of this range; however, the crude extracts (before sonication) used in the T15 assays had previously been frozen and stored. Beinor (1978) has reported that the freezing and storage of crude extracts can significantly decrease the activity for dehydrogenases. Also, the MDH activity is very sensitive to sonication conditions, as it is estimated that greater than 60% of the enzyme may be membrane-bound (Wadzinski and Ribbons, 1975a).
Undoubtedly, higher MDH activities for T15 would have been found with fresh extract preparations and optimiza¬ tion of sonication conditions; however, the primary purpose of the assay in this case was simply to demonstrate the presence of the enzyme.
iii) Glucose 6-Phosphate Dehydrogenase (GPP) and 6-
Phosphoqluconate Dehydrogenase (PGP)
Glucose 6-phosphate dehydrogenase is an NAD+ or
NADP+-linked enzyme found in the assimilatory pathway of all RMP bacteria. 6-phosphogluconate dehydrogenase also uses NAD+ or NADP+ as a coenzyme, and it catalyzes the key step in the cyclic oxidation pathway of RMP bacteria: the oxidative decarboxylation of 6-phosphogluconate to
ribulose 5-phosphate (refer to Figure 1). Therefore, 39
the presence of PGD in the cell-free extracts serves as a positive test for the involvement of the cyclic oxidation
route.
The average specific activities for T15 were as
follows:
Enzyme Specific Activity nmoles/min-mg Protein
GPD 111.2
PGD 50.6
Reported literature values for the two enzymes range
from 21 to 1580 nmoles/min-mg for GPD and 13 to 400
nmoles/min-mg for PGD (Steinbach et al., 1978; Ben-Bassat
and Goldberg, 1980; Anthony, 1982; Beardsmore et al.,
1982). Once again, the experimental activities for T15
were slightly low, probably due to the use of frozen
extracts for the assays. In any case, the presence of
these two enzymes confirms the involvement of both the
RMP assimilation pathway and the cyclic oxidation route
in the T15 strain. 40
III . T15 Batch Growth Experiments
A series of batch culture studies with T15 were performed in an effort to characterize the strain with respect to other methylotrophs. All batch cultures were incubated at 30°C.
i) pH Profile
As described earlier, the working liquid medium composition included a pH 6.9 phosphate buffer. In order to study the effect of pH on the specific growth rate of
T15, several different pH buffers were added to the liquid medium. In all cases, the buffers were 0.6M and were added at a concentration of 60 ml/1. The initial
MeOH concentration remained constant at 0.1% (v/v) in all of the cultures. Figure 2 gives the resulting pH profile. The T15 strain was capable of growth in the pH range studied, from 6.0 to 8.0. The highest growth rate
(u = 0.433 hr”^) resulted at a pH of 7.0. This was an expected result, since the bacteria had originally been isolated and selected on neutral media. Several methylotrophic species, such as Methylomonas EP-1,
Methylomonas BC-3, and Methylomonas L3 (Papoutsakis,
1976; Chen et al., 1977; and Hirt et al., 1978) have exhibited similar pH optima and specific growth rate profiles. SPECIFIC GROWTH RATE, hr" Figure 2.TheT15pH profile PH 41 42
ii) Growth on Pure Substrates
Methanol is the primary source of carbon and energy for bacteria which utilize the ribulose monophosphate pathway of carbon assimilation. Formaldehyde and formate are the partial oxidation products of MeOH, and they are key intermediates in the linear oxidation route of RMP bacteria for the production of biosynthetic energy, as mentioned earlier. Because of the vital role that these three substrates play in the metabolism and growth of RMP bacteria, batch growth studies on the pure substrates are of primary interest.
T15 growth on methanol alone was studied at different initial MeOH concentrations (SQ). Figure 3 shows the growth curves for S0 values of 0.2 and 2.0% (v/v). As the figure illustrates, T15 growth at the higher SQ resulted in a reduction of the growth rate from 0.32 to 0.22 hr”*.
This trend was observed in all of the batch cultures? in general, the lag phase increased and the specific growth rate decreased with increasing SQ. The T15 strain was able to growth at SQ values of up to 6% (v/v), but the calculation of growth rates in these cultures was diffi¬ cult due to the MeOH evaporation effects and prolonged lag phase (of the order of 5-7 days).
Fast serial transfers were performed with T15 in an effort to exercise the bacteria and select for natural OPTICAL DENSITY at 600nm Figure 3.Batchgrowth curvesforT15atdifferent initial methanolconcentrations 43 44 fast-growing mutants on pure MeOH. A batch liquid culture with 0.1% (v/v) MeOH was inoculated with T15, and as soon as growth in the flask was visible, a small inoculum was transferred to a new flask. Figure 4 shows the results of several of these serial transfers. The specific growth rate was improved from 0.32 to 0.43 hr“^ as a result of this natural selection process. The highest observed growth rate for T15 in any of the batch culture experiments was 0.46 hr-*.
Formaldehyde alone would not support the growth of
T15 at any of the concentrations studied (from 0.01 to
0.1% w/v). This was an expected result, as the majority of RMP bacteria have not been able to grow on formaldehyde as a sole carbon source (Anthony, 1982). However, a few
RMP species, such as Methylomonas L3 (Hirt et al., 1978) and Methylomonas EP-1 (Papoutsakis, 1976), did grow on formaldehyde at very low concentrations (<0.05% w/v), although the growth rates and biomass yields were very poor and the lag phases were very long.
The T15 strain would also not grow on formate as a sole carbon source. Initial formate concentrations ranging from 0.01 to 0.1% (w/v) were studied. This result is typical of RMP bacteria, because the branch point between oxidation and carbon asimilation in the RMP pathway is at the oxidation level of formaldehyde (refer OPTICAL DENSITY at 600nm 2.0 .03 .04 Figure 4. The effectoffast transfers onT15growth 3 456 TIME (hrs) 8 10 II 46 to Figure 1). Formate is simply oxidized to CC>2, and no route for carbon incorporation exists through formate or co2.
iii) Growth on Mixed Substrates
Batch growth studies of T15 on MeOH/formaldehyde and
MeOH/formate mixtures were performed in order to examine the inhibitory effects of the substrates.
Figure 5 shows the effect of formaldehyde on the specific growth rate of T15. The initial methanol concentration in these cultures remained constant at 0.1%
(v/v). As the figure shows, a formaldehyde concentra¬ tion of 0.04% (v/v) completely inhibited the growth in this mixed substrate culture; however, when exercised and subcultured for several weeks on high concentrations of formaldehyde, the T15 cells were able to grow at formal¬ dehyde concentrations in excess of 0.06% (v/v).
Methylomonas EP-1 (Papoutsakis, 1976) has exhibited an almost identical formaldehyde inhibition profile to the one diagrammed in Figure 5 for T15. A more detailed discussion of the mechanism of formaldehyde inhibition follows.
Mixed-substrate cultures of T15 on methanol (0.1% v/v) and formate were studied at formate concentrations ranging from 0.1 to 0.2% (v/v). The growth curves for SPECIFIC GROWTH RATE, hr’1 0.5 • 0 .01.02.03 .04 Figure 5.Theinhibitory effectofformaldehydeon T15 growth INITIAL CH0CONC(W/V) 2 47 48 both a control culture (MeOH only) and a culture con¬ taining 0.2% formate are shown in Figure 6. The growth curves (and hence the specific growth rates) were essentially identical. In fact, formate did not cause inhibition of T15 growth at any of the concentrations studied. This property of formate tolerance is apparently unique to T15, as formate is typically in¬ hibitory to the growth of RMP bacteria (Pilot and Prokop,
1975? Chen et al., 1977; Hirt et al., 1978).
iv) Discussion: The Mechanism of Substrate
Inhibition
At this point, a detailed discussion of the sub¬ strate inhibition effects of methanol, formaldehyde and formate is in order. Many empirical models have been proposed to describe inhibitory growth in methylotrophs
(Edwards, 1970; Wayman and Tseng, 1976; Chen et al.,
1976). For the most part, these empirical fits simply involved the modelling of the specific growth rate vs. substrate concentration curve. Also, simple enzyme- substrate inhibition kinetics have generally been assumed? that is, the rate controlling step was assumed to be a substrate-inhibited enzymatic reaction that occupied a key position in the carbon assimilation pathway
(Papoutsakis et al., 1978). In order to develop kinetic expressions, it was assumed that the intracellular OPTICAL DENSITY at 600nm Figure 6. Batch growthcurves withmethanolandformate 49 50 concentration of the enzyme which catalyzed the rate¬ controlling step remained constant. Clearly, such an assumption is not valid for most regulatory enzymes; in general, repression and induction mechanisms regulate the intracellular concentrations of key enzymes. Indeed, a number of researchers have found that the key enzymes in the biosynthetic pathways of methylotrophic bacteria were inducible (Goldberg and Mateles, 1975; Ben-Bassat and
Goldberg, 1980).
Experimental work by Papoutsakis (1976), Beinor
(1978), and Krug et al. (1979) has resulted in the development of a conceptual model for the inhibitory nature of compounds in RMP bacteria, based upon the induction-repression mechanism of enzyme formation. The key elements of the model are as follows:
1. Methanol is required for the induction of both
MDH and HPS synthesis.
2. High intracellular concentration of formaldehyde causes repression of HPS synthesis.
These conclusions provide an explanation for the inhibi¬ tion of growth at high MeOH concentrations, as observed with T15. The presence of MeOH within the cells induces the synthesis of the oxidative enzymes and HPS, and the methanol is subsequently oxidized to formaldehyde. Some 51 of this formaldehyde is oxidized to formate and CO2 via
the linear oxidation route (in order to provide biosynthe¬
tic energy), and some is incorporated into biomass via
HPS and the assimilatory pathway. As the concentration of methanol in the external medium is increased, the
intracellular concentrations of methanol and formaldehyde
also increase. A large internal formaldehyde concentra¬
tion represses the HPS synthesis, so that more of the
formaldehyde is oxidized via the linear route, and less
is assimilated into biomass. This results in lower
biomass yields and reduced specific growth rates.
Therefore, formaldehyde is the key intermediate for the
regulation of growth.
Formate also inhibits the growth of RMP bacteria,
although it is typically less toxic than formaldehyde.
For example, in Methylomonas BC-3, formate was about 10
times less inhibitory than formaldehyde (Chen et al.,
1977). The mechanism of formate inhibition is not known,
although it has been suggested that a feedback inhibition mechanism might be involved (Papoutsakis et al., 1978).
In the case of T15, the extremely high jin vitro levels of
formate dehydrogenase activity indicate that these cells
maintain high concentrations of FDH; consequently, the
intracellular formate pool is kept at a sufficiently low
level so that feedback inhibition is not a problem. This
explanation accounts for the high formate tolerance 52 exhibited by the T15 strain,
v) T15 Batch Yields
The biomass yield in batch culture is defined as follows:
r Y x/s x = dX/dt = dX (2) -rs -dS/dt -dS
where X = biomass concentration,
and S = substrate concentration
Integration of equation (1) gives
(3)
Rearranging,
X (4) Vs
since XQ = 0. Y x/s is defined as the average biomass yield over the course of growth of the batch culture (the superbar denotes average). Eq. (4) was used to calculate
T15 batch yields at initial methanol concentrations ranging from 0.025 to 3.25% (v/v). The results are diagrammed in Figure 7. Other microorganisms such as
Methylomonas EP-1 and Methylomonas L3 have exhibited a YIELD g cells/g MeOH 0.7 - 0 0.20.40.60.81.0 2.0 3.0 Figure 7. Biomass yieldasa function ofinitial methanol concentration frombatchexperiments INITIAL CHOH CONC.(%v/v) 3 53 54 similar dependence of the batch yield on MeOH concentra¬ tion (Papoutsakis et al., 1978; Hirt et al., 1978). One significant difference with the T15 strain, however, was
the exceptionally high yield at low MeOH concentrations.
For example, the measured biomass yield at 0.025% MeOH was 0.63 g cells/g MeOH. This value was higher than any other batch yield reported in the literature for methylotrophic bacteria (Papoutsakis et al., 1978; Anthony,
1982). In fact, the only microorganism reported to have
a higher yield under any conditions of growth was
Methylomonas EP-1, which exhibited a continuous culture yield of 0.65 (although the highest batch yield was only
about 0.35). Maximal yields for a particular strain are
typically achieved under steady-state chemostatic growth
(balanced growth), while growth in batch culture is
unbalanced, and as a result, batch yields are typically
lower. The fact that T15 exhibited batch yields in
excess of 60%, therefore, indicated that the micro¬
organism was extremely efficient in terms of carbon
assimilation, particularly at low MeOH concentrations.
As mentioned in the Introduction, computation of
biomass yields based upon theoretical considerations is
often valuable for the purpose of comparison with
experimental data. In this section, the method originally
developed by Van Dijken and Harder (1975) is used 55 to compute biomass yields for T15. The formula used to represent cell material in this development is C^gC^N, which is close to the composition measured for a number of methylotrophic bacteria (Maclennan et al., 1971, Goldberg et al., 1976). The linear pathway of methanol oxidation may be represented as follows:
CH2OH HCHO HCOOH ^ N ’ ^ V c°2 x XH2 Y YH2 z ZH2 where the letters X, Y and Z represent the electron acceptors involved in the oxidation steps. Numerical values for X, Y and Z represent the equivalent moles of
ATP formed during each oxidation step. In the ribulose monophosphate pathway (as well as the serine pathway), carbon is assimilated into biomass through the key
intermediate 3-phosphoglycerate (PGA); therefore, the calculation of the energy requirement for assimilation is divided into the following two parts:
1) Synthesis of PGA from units.
2) Synthesis of cell constituents from PGA.
Ths synthesis of PGA may be represented by the following
expression, based upon the reactions of the RMP cycle
(Kemp and Quayle, 1966; Strom et al., 1974):
3CH3OH PGA + 3XH2 + NADH2 (5) 56
The synthesis of cell material from PGA is represented by
Bq. 6:
4PGA + 3NH3 + 5.5 NADH2 + 29ATP ■ -» 3C4H802N (6)
The original paper (Van Dijken and Harder, 1975) presents
a detailed development of this expression. Elimination
of PGA and a combination of Eqs. 5 and 6 gives the
following overall expression for the synthesis of cell material from methanol:
12 CH3OH + 3NH3 + 29 ATP + 1.5 NADH2 3C4H802N + 12XH2(7)
Enzyme assays with the T15 strain proved that both
FDDH and FDH are NAD-1inked? Therefore, Y = Z = NAD and
Y = Z = 3, assuming that 3 moles of ATP are produced per
mole of NADH2 oxidized. Biomass yields for T15 may be
calculated from Eq. 7 by estimating the amount of methanol
required for the generation of NADH2 and ATP. Assuming
that X = 1, at least 0.75 moles of MeOH must be oxidized
in order to meet the NADH2 requirement, since NADH2 is
generated by the oxidation of both formaldehyde and
formate. Therefore, Eq. 7 becomes
12.75 CH3OH + 3NH3 + 29 ATP ^3 C4H802N + 12.75 X H2
where 12.75 XH2 corresponds to 12.75 moles ATP (since
X = 1). 57
Therefore, another 29 - 12.75= 16.25 moles of ATP must be formed from the oxidation of 16.25/1+3+3 = 2.32 moles MeOH.
T15 growth on methanol, therefore, is described by the overall equation
15.07 CH3OH + 3NH3 + 9.8602—►3C4H802N + 3.07CO2 +
22.64H20 (8)
The maximum yield in this case is given by
(3)(102 ) Yx/S = =0.63 g cells/g MeOH (15.07)(32)
Adding 3% to the theoretical yield to account for ash, as suggested by Van Dijken and Harder (1975) the maximum obtainable yield is 0.66. A similar development for X = 2 gives a maximum theoretical yield of 0.73. The experi¬ mental value of 0.63 for T15 shows that the yield has either closely approached the maximum theoretical yield in the case of X=l, or is only about 15% lower than the maximum value in the case of X = 2. These results, together with a comparison of T15 batch yields with those of other methylotrophic bacteria, suggest that T15 is indeed a bioenergetically superior species. The selection criteria based upon the high jji vitro activities of FDDH and FDH, threfore, has proven to be valid. 58
IV. Systematics: General Characterization of T15
A vast number of methylotrophic bacteria have been isolated and studied during the past 15 years, and this was primarily a result of the realization of the potential of these microorganisms for SCP production. Many of these bacteria were studied by researchers who had little interest or expertise in nomenclature and taxonomy; consequently, there has been little consistency in the classification of methylotrophs in the literature. Some researchers have adopted the nomenclature originated by
Whittenbury (1970 a,b), who was one of the first to isolate, study and characterize a large number of methylotrophic bacteria. Others have developed completely new names, or perhaps just a series of letters or numbers, to designate their strains.
Perhaps the most widely used classification scheme for bacteria is that given in Bergey's Manual of
Determinative Bacteriology (1974); however, for the case of methylotrophic bacteria, this publication is outdated and suffers from a lack of completeness. Anthony (1982), has suggested a classification scheme for methylotrophs based primarily upon the type of carbon assimilation pathway.
Rather than relying on the "authority" of any of these classification schemes, an attempt was made here to 59 further characterize the T15 strain by means of a variety of standard biochemical and microbiological tests. The purpose of these tests was not to assign an exact name to the T15 strain, but rather to provide comparative evidence, combined with the biochemical information and macroscopic growth characteristics described in previous sections, to characterize the strain with respect to other methylotrophic bacteria.
i) General Characterization
Nitrogen-purged, sealed fermentation tubes containing
0.2% (v/v) MeOH and the salts solution would not support the growth of T15. Similar sealed tubes containing air, however, did support growth. These results suggest classification of T15 as an obligate aerobe.
ii) Morphological Characteristics
Gram stain preparations of T15 have revealed that the strain is gram negative. The bacteria are long, straight rods with dimensions of 3 to 4 um by 0.5 to 1 um. Size estimations were made directly from formalin- fixed gram stain slides, or from photomicrographs of these slides. T15 was by far the largest of the three original soil isolates. Bacterial size is an important
industrial characteristic, since larger bacteria may be more efficiently and inexpensively harvested from the growth medium than smaller bacteria. 60
iii) Physiological Tests
The catalase and oxidase tests are commonly used for the general classification of a wide variety of bacteria.
The T15 strain exhibited a positive reaction to both of these tests.
The T15 strain did not grow on complex media
(nutrient agar), unless the media was supplemented with methanol.
T15 growth on a variety of carbon sources was studied using standard fermentation tubes. The results are given in Table 2.
Table 2: T15 Growth on Various Carbon Sources
Carbon Source Acid Production Growth Supported
Methanol + +
Formaldehyde + -
Formate - -
Ethanol + -
Glucose + -
Fructose + -
Sucrose + -
Acetate - -
Pyruvate + -
Glycine + - 61
Of all the carbon sources tested, only methanol was a suitable substrate for growth, as indicated by the turbidity in the tube. A subsequent study of T15 growth on these various carbon sources in standard batch culture flasks revealed that T15 was able to grow very slowly on glucose? however, the lag phase was long and the yield was poor. Othr species such as Methylomonas EP-1
(Papoutsakis, 1976) have exhibited similar growth on glucose. Although none of the substrates other than methanol was able to support efficient growth of T15,
Table 2 shows that T15 was able to oxidize these sub¬ strates, as indicated by the production of acid in the tubes. Incubation of T15 with formate and acetate resulted in the alkalinization of the medium, which was attributed to the uptake of the acids. This subject will be discussed in more detail later.
Based upon the results presented above, the T15 strain, being rod-shaped and utilizing the RMP pathway, is most similar to those methylotrophs which have been previously designated as Methylomonas or Methylobacter species. The genus Methylomonas, according to Bergey1s
Manual (1974), belongs to the Methylomonadaceae family.
The name Methylobacter finds its origin in the work of
Whittenbury et al. (1970 a,b), and this genus is not recognized by Bergey*s Manual. Methylotrophs belonging to other genera, such as Pseudomonas and Bacillus, also 62 utilize the RMP pathway; however, these bacteria are typically characterized by efficient growth on a wide variety of carbon sources (as well as on nutrient media), and they are not usually isolated on methanol (Anthony,
1982).
Many of the batch growth studies with T15, as described in earlier sections of this thesis, were patterned after studies with the Methylomonas species
EP-1, BC-3 and L3. Indeed, the results of these studies suggest that T15 is similar to these species in many respects (pH profile, methanol and formaldehyde inhibi¬ tion, etc.); however, a number of characteristics, such as size, formate tolerance and high batch yields dis¬ tinguish the T15 strain from these and other methylotrophic species. Exact classification of T15 into existing genera becomes more difficult, and to some extent less justi¬
fiable, due to these unique characteristics. Suffice it
to say that of all methylotrophs which have been charac¬
terized in the literature, T15 is most similar to the
Methylomonas and Methylobacter species, as mentioned earlier.
V. Continuous Culture Experiments with T15
Under steady-state conditions, the following mass
balance may be applied for any component in a chemostat 63 system:
Rate of addition to system - Rate of removal from system
+ Rate of production within system = 0
In terms of cell concentration, this becomes
F (xQ-x) + Vrx = 0 where F = volumetric flow rate of feed and effluent
streams,
x = viable cell concentration in reactor and exit
stream,
xQ = viable cell concentration in feed stream,
V = working volume in chemostat, and
rx = rate of cell formation
Let u = specific growth rate = —-—
and D = = dilution rate.
It follows that Dx0 = (D-u)x
For sterile feed, xQ = 0. Therefore,
0 = (D-u)x
or D = u
This development shows that under steady-state conditions 64 in a chemostat, the dilution rate is equal to the specific growth rate. A steady-state mass balance on the substrate (methanol) gives
F(S0-S) + Vrs = 0
ux _ _ Dx where r<= = rate of substrate utilization = - — Y Y r x/s x/s Dx Therefore D(S0-S) = ÿ x/s where Yx/s = g biomass formed/ g substrate consumed
Thus,
Yx/s - -s4l"
This equation is used to calculate steady-state biomass yields in a chemostat. The methanol feed concentration in all continuous culture experiments with T15 was 0.1% (v/v). Growth was studied at dilution rates ranging from 0.05 to 0.55 hr”l; however, all attempts to reach steady-state at dilution rates higher than about 0.1 hr--*- were not successful. Higher dilution rates resulted in washout of the cultures with high residual methanol concentrations. Several steady-states were established with T15 at or below dilution rates of 0.1 hr”l, and under these condi¬ tions the residual methanol concentration was essentially 65 zero. At steady-state, D = u; therefore, the highest observed specific growth rate for T15 in continuous culture was only about 0.1 hr-l. This value was significantly lower than the maximum observed growth rate for T15 in batch culture of 0.46 hr-^. This inability to grow at high dilution rates proved to be another unique property of the T15 strain. Others have reported speci¬ fic growth rates of at least 0.3 hr-1 for chemostatic growth of a variety of methylotrophic bacteria (MacLennan et al., 1971; Battat et al., 1974; Hirt et al., 1978;
Urakami and Komagata, 1979; Anthony, 1982). For example,
Methylomonas EP-1 was able to reach steady-states at dilution rates as high as 0.580 hr"-*- (Papoutsakis, 1976), and it is interesting to note that the growth conditions
(temp., pH, media composition, etc.) for EP-1 were essentially identical to those for T15.
Continuous culture yields for T15 were measured at several different reactor conditions (given in the
Appendix). The highest measured yield was 0.328 g cells/ g MeOH at a dilution rate of 0.101 hr--*-. This value was significantly lower than the highest batch yield of 0.63, and it was also much lower than continous culture yield values reported by other researchers. Methylomonas EP-1, for example, exhibited a maximum yield of 0.65
(Papoutsakis, 1976). 66
At first, it was suspected that the instability of
T15 cultures in the chemostat was resulting from inadequate reactor operating conditions (Temp, pH, agitation rate, etc.); however, operation of the chemostat in batch mode with identical conditions resulted in specific growth rates in excess of 0.40 hr“^. These results indicated that the conditions were adequate to support efficient growth of the strain. In addition, small adjustments to the operating variables at steady-state did not result in any significant improvement in the optical density of the culture, nor did these adjustments improve the ability of the strain to grow at higher dilution rates.
A media optimization procedure similar to the one developed by Matales and Battat (1974) was performed in order to examine the possibility that one of the nutrients was either limiting or inhibiting the growth of T15,
resulting in the observed instability of the cultures.
The initial liquid media composition for the continuous
culture was identical to the batch composition. After a
steady-state was achieved, each nutrient was aseptically
added to the reactor in order to give twice the original
concentration, and the response of the culture to each
pulse was observed. None of the pulses of the nutrient
salts caused any significant change in the optical density
of the culture, thereby indicating that these nutrients 67 were neither growth limiting nor inhibitory at the concentrations studied. When NH4SO4 was pulsed into the reactor to give three times the initial concentration, the optical density decreased and the culture began to
+ washout. These results suggested that NH4 was in¬ hibitory at very high concentrations. Others have observed inhibitory effects of NH4+ on methylotrophic growth (Whittenbury et al., 1970a), although the exact mechanism of this inhibition has not been established.
In any case, the concentration of NH4+ used in the continuous culture experiments with T15 was adequately low to ensure that inhibition was not a problem.
In experiments with L3, Krug et al. (1979) observed a buildup of formaldehyde in the chemostat medium during unstable growth and washout conditions. It was specu¬ lated the formaldehyde was building up within the L3 cells and inhibiting their growth via repression of HPS synthesis, resulting in the instability and washout of the culture. It seemed reasonable that the same mechanism might be responsible for the instability of
T15 growth at higher dilution rates. In order to test this hypothesis, samples were removed from the reactor under different growth conditions (steady-state, unstable growth, washout), filtered to remove the cells and preci¬ pitates, and assayed for formaldehyde by means of the 68 chromotropic acid method. Essentially no formaldehyde
(<0.2 mg/1) was found in any of these reactor samples.
These results suggested that simple substrate inhibition was not the cause of the reactor instability. In addi¬ tion, gas chromatography of the chemostat samples did not reveal any additional metabolites which might have been inhibitory to the growth of the strain.
Up to this point, then, no reasonable evidence had been found to explain the low washout dilution rate of
0.1 hr“l for T15. It was decided that the study of the substrate-transport mechanisms of T15 might help to explain some of the unusual growth characteristics of the strain, and as a result this avenue of study was pursued.
VI. Formate Uptake by T15t Proton Translocation
Measurements
When the T15 strain was grown on methanol or a combination of methanol and formaldehyde, the medium became more acidic as the growth of the culture proceeded.
For example, the initial pH of the buffered media
immediately after inoculation was usually about 7.0, and after the culture reached stationary phase the pH was
typically around 6.75. Much of this acidification may be attributed to the electron transport activity resulting
from the oxidation of the substrate via the cyclic and 69
linear oxidation routes. ATP hydrolysis through the
ATPase system, as predicted by the oxidative phosphyla-
tion theory of Mitchell (1970), would also account for a
release of protons from the cells resulting in a pH drop
of the external medium. It has also been speculated that
+ + the release of H from NH4 upon utilization of the
ammonia is responsible, at least in part, to the observed
pH drop during methylotrophic growth on methanol
(Papoutsakis, 1976).
When T15 cultures were grown on a combination of
methanol and formate, a profound alkalinization (rather
than acidification) of the medium was observed. For
example, a mixed-substrate culture with 0.1% (v/v) MeOH
and 0.1% (w/v) formate, which had an initial pH of 7.13
after inoculation, eventually reached a pH in excess of
7.7 at stationary phase. Alkalinization effects were
observed with formate concentrations as low as 0.01%
(w/v), although the final pH of the medium decreased with
decreasing formate concentration. In addition, pH
increases were observed when T15 cells were incubated in
media containing formate alone, although essentially no
growth was observed in these cultures. This showed that
the increase in pH was due to the prsence of the formate
rather than to some combined effect of the substrates.
It is interesting to note that the highest pH reached in
any of the cultures was about 7.75. Addition of more 70 formate to these cultures did not result in any additional increase in the pH.
Little attention has been paid to formate-induced alkalinization in the literature, although this effect has been observed with other methylotrophic bacteria
(Dijkhuizen et al., 1977b; Papoutsakis et al., 1981).
Two possible explanations exist for the observed pH increase with these cells:
1) The alkalinization is due in some way to the
oxidation of the formate.
2) The uptake or transport of the formate is
accompanied by co-transport of protons, result¬
ing in a pH increase of the external medium.
In RMP bacteria such as T15, oxidation of formate via formate dehydrogenase results in the production of NADH2 and CC>2. The NADH2 production will cause an increased H flow outwards due to the oxidation of the reduced coenzyme via the electron transport chain. This will result in acidification of the external medium. Furthermore, transport of CO2 out of bacterial cells is thought to involve a uniport system that is not associated with proton translocation (Mitchell, 1970). Also, oxidation of large quantities of formate occurs in methylotrophs when the cells are grown on methanol alone (or MeOH +
CH2O), and acidification rather than alkalinization is observed in this case, as mentioned earlier. It is highly 71
unlikely, in light of the reasons above, that the
alkalinization effect is due to the oxidation of the
formate. Therefore, the proton uptake must be due to the
transport of formate into the cells. Formate is an
anionic molecule which exists primarily in the form HCOO”
at the neutral pH of the growth medium. In order to
maintain electrical neutrality across the cell membrane,
the cells transport formate in the proton-coupled form
HCOOH. The symport of protons with organic acids is a
common mechanism in bacteria (Salanitro and Wegener,
1971; Matin and Konings, 1972; Gutowski and Rosenberg,
1975; Kell et al., 1981), and in particualr, the trans¬
port of formate across membranes as an undissociated
molecule has been observed by a number of researchers
(Dijkhuizen et al., 1977a; Garland et al., 1975;
Boonstra and Konings, 1977).
In order to study the formate-induced alkalinization
in a more controlled environment, proton translocation
studies were performed with aerobic T15 whole cell
suspensions. Figure 8 shows the rate of proton uptake
into T15 cells at different formate concentrations. The
initial pH of each suspension was adjusted to a common
value of 7.10 for these measurements. The proton uptake
in Figure 8 is expressed in terras of nanograms of protons
(H+ ions) per mintue per milligram dry weight of cells, 72 Ma ôUI-UJUJ/O *D Ou V
o z o o UJ H < 2 tr o u- of formate concentration Figure 8. Formate-induced proton uptake by T15 cells as a function
N to to io CM 0000000
+H MQ 6UJ.UIUI/4H UOI 6U J O 73 which is a common unit for proton translocation measure¬ ments. As the plot shows, the rate of proton uptake by
T15 cells was directly proportional to the formate concentration in the suspension. At formate concentrations above about 5mM, a saturation effect was observed because more formate was present in the suspension than the cells were able to utilize. Figure 8 also shows the rate of oxygen utilization in the suspensions, which provided a direct measurement of the rate of formate oxidation by the cells. The trend for the oxygen uptake rate followed the trend for the proton uptake rate almost exactly, indicating that the formate was immediately being oxidized upon transport into the cells. The high
in vitro activities for formate dehydrogenase observed with T15 cells would account for this rapid oxidation of
formate.
During the course of these experiments, it was
noticed that the rate of proton uptake by T15 cells was dependent upon the pH of the suspension. Figure 9 clearly
displays this phenomenon. The formate concentration
remained constant at 5.56 mM in these experiments. The
plot shows that as the pH of the suspension was adjusted
to more acidic values, the rate of proton uptake by the
cells increased. These results suggested that the formate
uptake by T15 was dependent upon the pH gradient across ng ion H*/min. mg DW Figure 9.ThepHdependence offormate-inducedproton andD =formate+CCCP. uptake. O=formate only,A=formate+KSCN, 74 75 the membrane. As the pH of the suspension was adjusted closer to the intracellular pH, the driving force for the formate uptake (ApH) was reduced, resulting in a decrease in the observed rate of proton uptake. Figure 8 also shows that when the cells were incubated with carbonyl cyanide m-chorophenyl hydrozone (CCCP final cone = 5 uM) prior to the formate injection, the rate of proton uptake was significantly reduced. CCCP is a proton ionophore which abolishes the ApH across cell membranes (Harold et al., 1974). These results further suggested the involve¬ ment of ApH with formate transport. Furthermore, when the cells were incubated with thiocyanate (KSCN final cone = 90 mM), which destroys the electrical potential across bacterial membranes (Mitchell and Moyle, 1969), the rate of proton uptake was not affected. Thus the
formate uptake did not appear to be dependent upon the AŸ component of the proton-motive force. These preliminary
results were later confirmed by the results of radio-
labelled formate uptake experiments. These experiments
are described in subsequent sections of this thesis.
One additional experimental observation deserves mention here. When formate was injected into anaerobic
T15 cell suspensions, the uptake of protons was still
observed. This provided yet another indication that 76 uptake, rather than oxidation, was responsible for the alkalinization of the medium.
VII. Radiolabelled Substrate-Transport Studies
The raw data and calculation procedure for the transport experiments with -^C-labelled substrates are given in the Appendix. Each experiment was repeated at
least once to ensure reproducibility of the data,
i) Measurement of T15 Cell Volume
For bacterial transport processes, it is often convenient to express uptake of a substrate in such a way which expresses the relationship between the extra¬ cellular and intracellular concentrations of the sub¬
strate, since it is this parameter which distinguishes between diffusion (no accumulation) and an active trans¬ port system (accumulation) (Maloney et al., 1975). In
order to measure the internal concentration of a substrate,
it is first necesary to estimate the intracellular volume.
This was accomplished for T15 cells by means of permeabil¬
ity measurements with l^C-labelled glycerol and sucrose
(Palmieri and Klingenberg, 1979; Marquis, R.E., 1981).
Glycerol readily permeates cells, but is not concentrated
by them, while sucrose is permeable into the periplasmic
space of gram-negative bacteria, but does not penetrate
the inner cell membrane. 77
The procedure for these labelled permeability measurements was the same as for the other labelled
experiments, except that the glycerol and sucrose were
allowed to equilibrate for much longer times (from 10-20
min), and samples were extracted from the suspensions
less frequently. The permeability profiles for glycerol
and sucrose are shown in Figures 10 and 11, respectively.
Figure 10 displays the gradual equilibration of glycerol
across the cell membrane. The profile for sucrose, on
the other hand, showed an initial peak of labelled
activity in the cells, followed by a gradual release and
equilibration of the sucrose. Absorption of the sucrose
on the membrane surface might have been responsible for
the initial peak. In any case, the regions of interest
in these plots were at higher equilibration times when
the labelled activity in the cells had reached a constant
value. The ratio of radioactivity in the cells to that
in the supernatant was used to compute the cell volume
(calculation procedure in the Appendix). These experi¬
ments gave a total cell volume of 8.0 ul/mg DW. Almost
1/5 of this volume was sucrose-permeable, corresponding
to 1.5 ul/mg DW of periplasmic space. These results
were very close to the values found for Methylomonas L3
by Diwan et al. (1983). 5000 ST130 NI INdQ aaznvwaoN
Figure 10. Glycerol uptake profile 78 12000
100 200 300 400 500 600 700 800 900 1000 TIME (sec) Figure 11. Sucrose uptake profile 80
ii) l^C-labelled Methanol Uptake
The uptake profile for methanol is shown in Figure
12. The external concentration of methanol in these experiments was 77 uM. Experiments with other methanol concentrations resulted in similar uptake profiles. The
arrow on the right-hand side of the plot represents the
level of intracellular substrate (nmol/mg DW) that
corresponds to an equimolar concentration of methanol in
the cells and in the supernatant. Any value above this
level represents accumulation of methanol in the cells.
As Figure 12 indicates, the methanol eventually equili¬
brated to give a nearly equimolar concentration in the
cells and the supernatant. The profile, however, was
not a simple diffusion pattern (as with glycerol for
instance), but it displayed an initial peak which
corresponded to about a 2-fold accumulation of methanol
in the cells. The fact that the concentration eventually
equilibrates across the cell membrane, however, suggests
that methanol transport by T15 cells is a diffusion-
dominated process. This subject is discussed in more
detail later.
iii) l^C-labelled Formaldehyde Uptake
The external concentration of formaldehyde in these
experiments was 73 uM. The uptake profile is given in 14 Figure 13. The pulse of CH2O led to an initial I Figure 12. Methanol uptake profile
COCD^rOJOCOCO^J-CJ • » • • • • • • • — — — — — oooo e Ma BUI/IOUIU S||30 ui HO HO 82 accumulation of the substrate corresponding to a 6.5-fold concentration gradient. This initial peak was followed by an efflux of formaldehyde from the cells, due to the high permeability of the molecule. During this efflux phase, it has been speculated that some electron trans¬ port activity takes place, leading to the production of
ATP which drives the next phase of active transport
(Papoutsakis, et. al., 1983). Figure 13 shows this second phase clearly, as formaldehyde was again accumulated in the cells to nearly the same level as in the initial phase.
The second active transport phase was again followed by a diffusion-dominated efflux of the accumulated formaldehyde.
Incubation of the cells with Flouryl Cyanide m-Chlorophenyl
Hydrazone (FCCP final concentration = 5uM) prior to the
■*■ 01120 pulse completely abolished any significant accumulation of formaldehyde by the cells. FCCP is a proton ionophore which is similar in structure and function to CCCP, but is more potent. Incubation of the cells with thiocyanate (KSCN final concentration = 90 mM) did not affect the uptake profile significantly. These results suggested that the ApH component of the proton- motive force, rather than the Ai» component, was the driving force for the accumulation of formaldehyde by these cells. Involvement of the ApH and accumulation of substrate are indicative of an active transport system. O 83 CO
o *•
o CM
O O
o o> <0
S tii
O CO
O Figure 13. Formaldehyde uptake profile
O CM 84
iv) l^C-labelled Formate Uptake
For the most part, the data from the methanol and
formaldehyde uptake experiments were fairly reproducible, without any significant variation from experiment to experiment; however, the data from the labelled formate uptake experiments were much less consistent. Although all of the profiles showed the cycling of the formate molecule into and out of the cells, the results did not
follow a reproducible pattern (as with formaldehyde uptake). Figure 12 gives the uptake profiles for one
series of experiments. The formate concentration in
these experiments was 23 uM. In reviewing these plots,
it is most useful to observe the overall levels of
substrate accumulation under the different experimental
conditions. The formate uptake displayed peaks cor¬
responding to 5-fold accumulation of formate in the
cells. Incubation of the cells with KSCN (final concen¬
tration = 90 mM) did not effect the overall level of
formate accumulation significantly, and some of the
samples showed nearly a 5-fold concentration gradient.
Incubation of the cells with FCCP (final concentration =
5uM), however, resulted in very little accumulation of
the formate. These results were in agreement with the
preliminary proton translocation measurements described
earlier; that is, they suggested that the formate transport 85 Figure 14. Formate uptake profile 86 was driven by the ApH component of the proton-motive force.
The A¥ component was apparently not involved in the
transport process, based on the measurements with
thiocyanate. Accumulation of substrate and the involve¬ ment of the A pH are indicative of active transport.
v) Discussion
Transport experiments with small molecules such as
compounds are more complex than studies with larger molecules such as carbohydrates or amino acids, due to
the high diffusivity of the small molecules through cell membranes. Diffusion, therefore, plays an important role
in the .uptake of small molecules, even if a specific
active transport system exists for the molecule. In this
case, the overall transport is accomplished through a
combination of the diffusion and active transport
mechanisms.
The observed methanol uptake profile with T15 cells
is not a simple diffusion pattern; instead, the profile
shows reproducible peaks which correspond to a small (2-
fold) accumulation of the substrate. These results
suggest the possible presence of a weak active transport
mechanism for methanol in these cells. It is possible
that the energy-coupling mechanism for this transport is
inefficient, resulting in only small levels of accumula¬
tion. Alternatively, the active transport process for 87 methanol in this strain may have been reduced as the
result of natural mutation. It is also possible that
small amounts of methanol are transported actively by the
permease (membrane transport protein) for formaldehyde,
since the two substrates are very similar in structure,
particularly in aqueous solution. Although these are
just speculations, the fact remains that methanol
equilibrates to nearly equimolar concentrations during
uptake studies with T15, indicating that diffusion is
the dominant process for methanol transport by these
cells. By comparison, methanol transport into Methylomonas
L3 cells under identical assay conditions results in
concentration gradients from 6.5 to 15-fold (Diwan et
al., 1933). In addition, the methanol uptake profile
for L3 shows a distinct biphasic pattern, similar to the
one exhibited for formaldehyde uptake in T15, and no
equlibration of the methanol is observed with these
cells.
The results of the methanol uptake studies with T15
provide an explanation for the growth instability of the
strain in continuous culture. It has been suggested
that in order to maintain the high washout dilution
rates observed in continuous culture for some species,
it is necessary for the cells to maintain high intracellular
concentrations of methanol (Diwan, 1983). Indeed, steady- 88
States were achieved at dilution rates in excess of 0.58 hr“l with Methylomonas EP-1, although the external concentration of methanol was essentially zero (Papoutsakis,
1976). Methylomonas L3 also has exhibited a relatively high washout dilution rate (in excess of 0.5 hr“M in chemostat cultures (Krug et al., 1979).
Methanol transport studies with L3 also have shown that these cells accumulate methanol via an active transport mechanism, as mentioned earlier. T15, on the other hand, with its diffusion-dominated methanol uptake mechanism, is not able to accumulate methanol in suffi¬ cient quantities to support growth at high dilution rates. As a result, the cultures wash out at dilution rates above 0.1 hr“l. This also accounts for the absence of formaldehyde in the T15 chemostat samples, since substrate inhibition is not the cause of the wash¬ out; instead, the inability of the organism to accumulate methanol results in the instability of the cultures.
In terms of SCP production, the inability of T15 to grow at high dilution rates in the chemostat is not an attractive property, due to the low reactor productivity.
The T15 strain, however, might be best suited for a fed batch reactor operation. In a fed batch reactor, the concentration of methanol could be kept low at all times in order to take advantage of the very efficient growth 89 of the microorganism at low MeOH concentrations, as sug¬ gested by the T15 batch yield data. It is interesting to note that the lack of an efficient active transport system in T15 may account in part for the high batch yields of the strain, since energy expenditure is not
required for transport of the substrate.
Transport of formaldehyde by T15 cells showed a
nearly identical pattern to that found with L3 cells
(Diwan et al., 1983). In both cases, the uptake profile was biphasic, and the accumulation was dependent upon the membrane A pH. Measured accumulation levels of formalde¬
hyde in L3 cells were 10-fold, which were slightly higher
than the 6.5-fold gradients measured in T15. These
levels of accumulation may at first seem small, parti¬
cularly in comparison with the high accumulation levels
(in excess of 100-fold) which are commonly generated by
bacterial active transport systems. It must be remembered,
however, that formaldehyde is highly toxic to the growth
of methylotrophic bacteria. It would not be desirable
for the cells to accumulate large intracellular concentra¬
tions of formaldehyde, as this would lead to the inhibi¬
tion of growth through the repression of HPS synthesis.
In addition, formaldehyde is highly permeable, as noted 90 earlier, and efflux of formaldehyde from the cells by diffusion results in lowering the overall accumulation
level.
The primary function of the active formaldehyde
transport system during growth on methanol might be to
counteract the leakage of formaldehyde from the cells.
Upon production of formaldehyde from methanol by the membrane-bound methanol dehydrogenase, molecules of
formaldehyde inevitably leak across the cell membrane.
Rather than relying on a simple diffusion mechanism to
regulate the flow of formaldehyde back into the cells, it
is possible that these bacteria have developed active
transport systems for this purpose*in order to minimize
the loss of formaldehyde to the external medium. The
decreased loss of formaldehyde due to leakage would
result in a greater availability of formaldehyde for
oxidation and assimilation into biomass, which would, in
turn, lead to more efficient growth and higher biomass
yields.
The transport of formate by T15 cells proved to be
more intriguing, as well as more difficult to study. The
14C tracer expeiments were much less reproducible with
formate than with methanol or formaldehyde, although the
results generally displayed the cycling of the formate
into and out of the cells. The transport was 91 found to be dependent on the membrane A pH, and accumula¬ tion levels of about 5-fold were measured in the cells.
One possible explanation for the observed variation in the formate uptake data deals with the effect that organic acids have on the permeability properties of cell membranes. Certain organic acids are known to behave as uncouplers which allow protons to enter the cell from the external medium, thus destroying the A pH gradient (Foster and McLaughlin, 1974). Herrero (1983) has provided a detailed explanation of the uncoupling mechanism. Apparently, the unionized species of the acid partitions in the cell membrane, and the internal and external concentrations equilibrate. Because the intra¬ cellular pH is higher than the pH of the external medium, the dissociation of the acid proceeds to a greater extent inside the cell. This results in a net efflux of acid anions due the higher concentration of anions in the cell cytoplasm. It is believed that the exit of the charged anion from within the cell occurs when the anion collides with an undissociated molecule partitioned in the membrane. This collision gives rise to a dimer
(HA2”). Since the species which permeates the cell is the undissociated acid (HA) and the species leaving the cell is the dimer (HA2”), for every molecule of acid entering the cell one proton is internalized. The 92
internal concentration of the acid anion is subsequently
replenished by diffusion of undissociated molecules from
the external medium. The uncoupling mechanism, then,
results in a cycling of the anion across the cell membrane.
Such a model has been used to successfully predict growth
inhibition by acetic, propionic, butyric and 0-hydroxy-
butyric acids (Herrero, 1983). This type of model would
account for the random cycling of the formate anion
observed in formate transport studies with T15. This model would also account for the proton uptake accompany¬
ing the formate transport. It is quite possible that
the transport of formate occurs by a combination of an
active transport system, which would account for the
observed accumulation, and an uncoupling mechanism. It
is interesting to note that incubation of T15 in batch
cultures containing acetate as a sole carbon source also
resulted in the alkalinization of the external medium.
This suggests that T15 might transport formate and acetate
(and possibly other acids) by a common transport mechanism. 93
CONCLUSIONS AND RECOMMENDATIONS FOR FUTURE WORK
From the results and trends observed in the previous section, the following conclusions can be drawn concerning the T15 strain:
1. T15 is an obligate methylotroph which utilizes the
ribulose monophosphate pathway of carbon assimila¬
tion .
2. The selection criteria based upon the jji vitro
activities of formaldehyde and formate dehydro¬
genase are valid. The T15 strain exhibits biomass
yields of up to 0.63 in batch culture, which are
higher than any batch yields reported in the
literature for methylotrophic bacteria. The
strain is particularly efficient at low methanol
concentrations.
3. The formate tolerance exhibited by T15 is unique
to this strain, and may be attributed to the very
high levels of formate dehydrogenase maintained by
the cells. This enzyme serves to rapidly reduce
the intracellular formate pool and reduce the
possibility of inhibition by this substrate.
4. Methanol enters the T15 cells primarily by means
of a diffusion mechanism. This accounts for the 94
relatively low washout dilution rates observed
in chemostat cultures, as the cells are not able
to accumulate methanol at high enough levels to
support efficient growth at high dilution rates.
5. Formaldehye is transported by T15 via an active
transport system which is driven by the A pH
component of the proton-motive force. The
membrane electrical potential (A'P) does not
appear to be involved in the transport. These
results are in agreement with experiments for
Methylomonas L3.
6. T15 cells have the ability to accumulate formate
by the membrane ApH, but the system is complex,
possible due to the uncoupling effect of formate
on the cell membrane. Formate uptake is
accompanied by the co-transport of protons,
which accounts for the alkalinization of the
medium during growth of the strain in mixed-
substrate cultures of methanol and formate.
Several of the properties of T15 suggest the need for further work. First of all, the growth of T15 should be studied in different reactor designs, such as fed batch, in order to take advantage of the superior effi¬ ciency of the strain at low methanol concentrations.
Based upon the work with T15 presented here, and in view of previous experiments with other RMP bacteria, 95 it is highly probable that a species exists which has high in vitro activities for FDDH and FDH as well as an efficient active transport system (permease) for methanol uptake.
It would be possible to select for such a species on the basis of enzyme assays and chemostat growth studies (and/or transport assays). A species with these characteristics might prove to be superior for industrial applications, since it would have both a high energy-producing capability
(due to the high FDDH and FDH activities) as well as the capability to grow efficiently at high dilution rates in continuous culture.
It is also recommended that the possibility of induction and repression of the substrate uptake mechan¬ isms be studied. This could be accomplished by assaying the activities of the transport systems under different growth conditions. For example, the uptake could be assayed at the different stages of growth in batch culture, or with different extracellular concentrations of the key substrates. Also a comparison of the transport activities in batch and continuously grown cells might be interesting. Studies such as these should help to elucidate the mechanisms which regulate the flow of the carbon substrates into the cells. These mechanisms are extremely important, as they have a profound influence on the overall biomass yield. 96
The energy coupling mechanisms of the T15 formal¬ dehyde and formate transport systems still need to be examined. Labelled uptake studies with arsenate, similar to those performed with L3 (Diwan et al., 1983) might provide valuable insight into the energy requirements of these transport processes. It is also suggested that uptake experiments with thiol reagents be performed in order to help determine whether or not a permease actually exists for methanol transport in T15.
The presence of active transport mechanisms in methylotrophic bacteria implies that simple unstructured models are not adequate for process control, since the substrate transport plays such an important role in the dynamics of the cell growth. More sophisticated models, which incorporate mathematical expressions for the substrate transport mechanisms, need to be developed for the purpose of fermentation control, as suggestd by
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Zatman, L. J. (1981) Microbial Growth on Ci Compounds (H. Dalton, ed.) Heydon, London. APPENDIX TABLE Al: Formaldehyde DH Activities for Isolated Strains S w •H •H 4-1 C . I ÿ s& •H 8 £ CÆ f-t t< £2 H O P •iJ •H 0 £ £ 8® O g fj-sl & O S 4J ta S1 (0 U r 4-) -H > <|J3 U Ü)
Table A3: T15 3-Hexulcse Phosphate Synthase Activities
Reaction Time Initial Œ2O Final CH2O Specific Activity /<• moles/tube /i moles/tube /L moles/hr’mg protein
1 27.31 11.53 383.5] 2 27.31 6.88 228.8V Avg = 263.9 3 27.31 5.39 179.2J 5 27.31 3.58 118.9 10 27.31 2.22 73.7 10* 22.31 1.08 86.5 10** 22.31 3.72 67.6
*Extract prepared 6-29-83; Protein Cone = 4.908 mg/ml **Extract prepared 7-21-83; Protein Cone = 5.503 mg/ml For all other assays, Extract prepared 8-4-84; Protein Cone = 6.013 mg/ml 105
Figure Al. HPS activity as a function of reaction time 106
VO LO • 00 II
rH (0 £ 2 LH ID rH X • • to w O O 4J O 4-t H O 00 CO o o ON in • • m £ 10 •H M< ^ •• CO CO 00 00 ON to CM I n* 10 r- o o • • CO CO ■8 U ? to & Cu B Û4 to -p 4J -H 2 i §s * I -3 Table A6: T15 6-Phosphoqluconate Dehydrogenase Activities S. 03 S’ rH X £ H O •H (D o o S. s? H S’ C0 O <4H (d &3 r a t H 4J -H > rH CO m ,pe <0 4J o w t § t a> Cil c i' C co m m r- o m tr> o a\ m 00 CO CO co O rH «* co CM r- CM • • • • O VD m il m co * * CM r- CM 0> O GO O O <0 O m c d o o (O io o CM Figure A2. T15 cell dry weight determination O Table A7 T15 Batch Yield Data Yield [Œ3PH]i Avg OD q cells mg CH^OH utilized q cells % v/v @ 600 rm L ' g CH3OH 0.025 0.286 0.130 205 0.634 0.065 0.566 0.266 515 0.517 0.100 0.416 (2/1)* 0.385 795 0.484 0.125 0.439 (2/1) 0.408 975 0.419 0.230 0.633 (2/1) 0.596 1830 0.326 0.340 0.660 (2/1) 0.623 2255 0.276 0.520 0.810 (2/1) 0.768 3880 0.198 0.750 0.574 (3/1) 0.808 3415 0.237 1.790 0.628 (3/1) 0.887 7740 0.115 1.950 0.616 (3/1) 0.870 7020 0.124 3.250 0.707 (3A) 1.002 21700 0.046 *Denotes dilution of CD sample TABLE A8: Continuous Culture Yields for T15 w •HI rH rH 10 ÏM 0) P2 C ■ol +j -H •H S' CO Ç? O rH •H g at U g 4J (D'd O 4-> Calculation Procedure for Substrate-Transport Experiments A. Proton Translocation Measurements Proton Uptake: The pH scale was calibrated by the injection of 10 ul of 5 mN HCL into the suspension. 10 ul of 5 mN HCL contains 50.362 ng ion H+. Rate of proton uptake by T15 = Rate of proton uptake after formate injection - Rate of proton translocation by endogenous substrate Each rate is given by the following: + A^mm x chart speed (cm) x 10mm x 50.362 ng ion H Ax (min) cm Ay from pH calibration mm mm r + x 1 _ ng ion H mg DW in suspension min-mg DW Oxygen uptake: The oxygen scale was calibrated so that Ay = 200mm at a setting of 100 millivolts full scale (mv/fs). Literature value for solubility of oxygen in KCL at 33°C = 4030 ng at 0/ 10ml. Rate of oxygen uptake by T15 = Rate of oxygen uptake after formate injection - Rate of oxygen uptake by endogenous substrate. Each rate is given by the equation on the following page 113 Ay.mm ^ chart speed (cm) ^ Suspension volume (ml) ^ 10 mm Ax (min) mg DW in suspension cm mm scale °2 sensitivity (mv/fs) 4050 ng at 0 100 mv/fs 10 ml 1 ng at 0 X 200 mm min-mg DW B. Labelled Substrate Uptake Experiments The specific activities of the labelled substrates used in the transport assays were as follows: ^CHjOH = 0.1684 pmole/dpm ■^CI^O = 0.0527 pmole/dpm 14CH00H = 8.675 X 10"3 pmole/dpm ES = external standard ratio (from Beckmann counter) The experimental correlation for the counting efficiency curve of the Beckmann Scintillation Counter was as follows: Counting Efficiency = (1.77)(ES) - 0.893 dpm = cpm/ Counting Efficiency = dpm values given in Tables A9 through A23 Blank correction = Correction for backround radioactivity = 60 dpm 114 dpm^ = dpm - 60 Total dpm for each sample = dpm^ for supernatant + dpm^ for filter samples = td In order to account for pipetting errors in the experiments, and in order to put the data from each experiment on the same basis, the dpm values were normalized to a common total value, tn< Typically, this value was the average of the total counts in a series of experiments. The normalized dpm values were then given by the following expression: dpmn = (dpmb)(tn/td) The retention of radioactivity due to the binding of the labelled substrates on the membrane filters was estimated at 3% of the total counts, based upon experimental measurements by Anil Diwan. The normalized dpm values (corrected for filter retention) given in Tables A9 through A23 were given by the following expressions: Normalized filter dpm = dpmn filter - (0.03)(tn) Normalized wash dpm = dpmn wash + (0.03)(tR) Cell and periplasmic space volume estimates: Each sample removed from the suspension was 50 ul total. 115 The ratio of the normalized dpm (corrected for filter retention) in the filters to the normalized dpm in the supernatant, after the equilibration of the labelled glycerol and sucrose, was used to calculate the desired volume as follows: Normalized filter dpm Cell Volume Normalized wash dpm X Supernatant Volume (from Glycerol experiment) Periplasmic Volume = Normalized filter dpm Normalized wash dpm X Supernatant Volume (from Sucrose experiment) Substrate Concentration in Cells: Total sample volume = 50 ul The normalized concentration of the labelled substrate in the T15 cells was given by the expression on the next page. 116 Normalized filter dpm X Specific activity of substrate ^ 1 ^ 1 ^ 1000 ul ^ nmole cone of susp (mg DW/ml) 50 ul ml 1000 pmole _ nmole mg DW TABLE A9: Glycerol Uptake Experiment #1 •H rH x: a 4-> *11 w u <1> to □ ^ ? to 0 I *o < o VO 00 vo m VO in CNH^iovoHCO^H^roroiO^rovoiOmiOoo oooooooooooooooooooo VO VO ON r* co r-r^r-r^r-r^r^r-r-r-voiovovovovovovovovo co in r- m oooooooooooooooooooo O orHOOOONONO>OSO>O\ON0NOON r* ON rH HHOOHHHOHO^^O>00(Tl^^ai^ • •••••••••••*••••••• VO VO 00 cn CN HHcNroin^h'Oocri iHiHcNoomior-oocrv çj*csa\^^(Nooo^r^^cM • ••••••••••••••••••a • rHrH*HrH H O OCOOOOOOOrHrH ONfOCOHCMHino^DCO • ••••••••••••••••••• i—J —I __Ji_4«I >IiH*4»4«I I oooLTio^^mro^hCN oooooooopo r-HHrorocs noomoo r^inovornmocMOCM houomroo^cNOm (NrfiH^fVOoOOinCTSi—I 04 HCMrH inmvOHOOoroh^oh CM VO^00 • ••••••••• I Hinroo I VO in r- •H •H i u s -M II a o gl or a