GLUCONATE METABOLISM IN LACTOBACILLUS AND ITS ROLE IN PERSISTENCE IN THE HUMAN INTESTINE
DISSERTATION
Presented in Partial Fulfillment of the Requirements for
the Degree Doctor of Philosophy in the Graduate
School of The Ohio State University
By
Julie Kay Jenkins, M.S.
* * * * *
The Ohio State University 2005
Dissertation Committee:
Dr. Ahmed E. Yousef, Adviser Approved By
Dr. W. J. Harper
Dr. Luis Rodriguez-Saona ______Adviser Dr. Polly D. Courtney Graduate Program in Food Science and Nutrition ABSTRACT
The demand for functional foods that provide a health benefit beyond basic
nutrition is on the rise. Prebiotics are non-viable food components that promote
the growth of a number of beneficial bacteria that naturally inhabit the human
intestinal tract. Some of these beneficial bacteria, such as Lactobacillus and
Bifidobacterium species, are capable of eliciting health benefits such as reduced
incidence of lactose intolerance and diarrhea, improved immune response and
reduced cancer risk. The present studies address the effect of gluconate, a
component of many foods and food supplements, on human intestinal
microflora. Genes involved in gluconate metabolism in Lactobacillus reuteri were characterized.
The effect of dietary gluconate supplementation on human intestinal populations of Lactobacillus, Bifidobacterium, Propionibacterium, and Escherichia coli
was assessed over a 7-week period. Weekly stool samples were collected from 12
subjects and selected bacterial populations were enumerated. Colonies were
subsequently grown on medium containing glucose or gluconate as the primary
carbon source. At the level tested, dietary calcium gluconate did not
significantly affect the overall quantity of Lactobacillus, Bifidobacterium, E. coli or ii Propionibacterium in most of the subjects tested. However, the proportion of gluconate-fermenting Lactobacillus and Propionibacterium strains increased upon
gluconate consumption in some subjects, suggesting that a shift in species
composition may occur. Speciation using the 16S-23S rRNA intergenic spacer
region showed 2-6 different gluconate-fermenting Lactobacillus species in each
subject and 11 different species overall.
A 500 base pair (bp) putative gluconate kinase and a 750 bp putative
gluconate permease fragments were found in the Lactobacillus reuteri 100-23
genome by amplifying DNA with degenerate primers. A gluconate-negative
mutant of L. reuteri 100-23 was constructed via homologous recombination
utilizing the putative gluconate permease fragment. The mutant (100-23D) and
wild-type strains were characterized for gluconokinase, 6-phosphogluconate
dehyrogenase and gluconate uptake activities. The mutant had significantly
lower gluconokinase and gluconate uptake activities compared to wild-type.
This suggests that gluconate genes in L. reuteri 100-23 are induced by gluconate
and potentially arranged in an operon. Transcriptional analysis of wild-type and
mutant strains showed a single transcript, indicating the gluconate permease
gene is constitutively transcribed, and not inducible, in the presence of glucose or
gluconate. Knowledge of gluconate fermentation in Lactobacillus will aid
researchers in their understanding of probiotic organisms, potential prebiotic
compounds and the natural human intestinal microflora.
iii
Dedicated to my family, especially my late Grandma Dee and Papa, whose
encouragement led me to further my academic career
iv ACKNOWLEDGMENTS
I would like to thank Dr. Polly Courtney, for her encouragement, direction and advice. I would like to also thank Dr. Ahmed Yousef for serving as my major adviser in Dr. Courtney’s absence. Thank you to Dr. Harper for his advice and encouragement. Thanks to the Ohio Agricultural and Development Center and the Swiss cheese consortium for funding this research.
I would also like to thank my friends, Nurdan Kocaoglu-Vurma, Corunda
Pruitt, Karen Flinger, Aaron Malone, Joy Waite and Olga Anggraeni for their emotional and technical support. I would also like to thank Rosemary Hage, who offered countless hours of advice. Most of all, I would like to thank my husband and my daughter for their never-ending patience, tolerance and encouragement.
v VITA
January 13, 1975...... Born, Rochester Hills, Michigan
May 10, 1997...... B.S. Biology/B.A. Chemistry Olivet Nazarene University Kankakee, Illinois
June 2000...... M.S. Food Science and Nutrition The Ohio State University Columbus, Ohio
June 2000- present ...... Research Associate Food Science and Technology The Ohio State University Columbus, Ohio
PUBLICATIONS
Research Publications
1. Jenkins, J.K. and P.D. Courtney. 2003. Lactobacillus growth and membrane composition in the presence of linoleic or conjugated linoleic acid. Canadian Journal of Microbiology. 49:51-57.
2. Jenkins, J.K., W.J. Harper, and P.D. Courtney. 2002. Genetic diversity in Swiss cheese starter cultures assessed by pulsed field gel electrophoresis and arbitrarily-primed PCR. Letters in Applied Microbiology. 35:423-427
FIELDS OF STUDY
Major Field: Food Science and Nutrition
vi TABLE OF CONTENTS
Page
Abstract...... ii
Dedication ...... iv
Acknowledgment...... v
Vita ...... vi
List of Tables...... ix
List of Figures ...... x
Chapters:
1. Literature Review...... 1
1.1 Bacterial gluconate transport and catabolism...... 2
1.2 Human intestinal microflora ...... 5
1.3 Role of E. coli gluconate metabolism and colonization of mouse intestine ...... 7
1.4 The effect of dietary components on intestinal microflora .8
1.5 Gluconate as a potential prebiotic dietary component...... 9
1.6 Molecular biology approaches to assess complex bacterial communities ...... 10
1.7 Rationale and significance ...... 12
vii
1.8 Hypothesis and goals of project...... 13
2. The effect of dietary gluconate on fecal content of human Lactobacillus, Bifidobacterium, Propionibacterium and Escherichia Isolates ...... 14
2.1 Abstract...... 14
2.2 Introduction ...... 15
2.3 Materials and methods...... 17
2.4 Results...... 22
2.5 Discussion ...... 25
3. Genetic characterization of gluconate genes in Lactobacillus reuteri 100-23 ...... 38
3.1 Abstract...... 38
3.2 Introduction ...... 39
3.3 Methods...... 41
3.4 Results and discussion ...... 61
Conclusion ...... 70
Appendix A: Evaluation of a Swiss cheese model system ...... 72 A.1 Abstract ...... 73 A.2 Introduction ...... 73 A.3 Materials and methods...... 74 A.4 Results...... 80 A.5 Discussion ...... 81
List of references ...... 95
viii LIST OF TABLES
Table Page
2.1 Media and growth conditions used to enumerate bacterial cell counts and percent gluconate fermenters in fecal samples……………………………………………………………………...... 20
2.2 Percent gluconate-fermenting propionibacteria pre-, during and post-gluconate treatments……………………...………………………...24
2.3 Lactobacillus isolates determined by sequencing the 16S-23S rRNA intergenic region……………………..…………………...29
3.1 Amino acid sequences of degenerate primer sets used to search for L. reuteri 100-23 gluconate genes…….……..…………...44
3.2 Growth of selected Lactobacillus, Bifidobacterium and Propionibacterium strains from human or food origins with gluconate or glucose as the sole carbon source………………………….…………………………..62
3.3 Enzyme activities of L. reuteri 100-23 and 100-23D…………………….……67
3.4 Gluconate uptake rates for L. reuteri 100-23 and 100-23D……………….….67
A.1 Culture combination recommendations from four different culture suppliers used in the manufacture of Swiss cheese and Swiss slurries……………………………75
ix LIST OF FIGURES
Figure Page
1.1 Entner-Doudoroff pathway………………………………………………..…...3
2.1 Calcium supplementation and sampling time line……...……………...... 18
2.2 Lactobacillus cell numbers pre-, during and post-gluconate treatments ……………………………………………….…...32
2.3 Bifidobacterium cell numbers pred-, during and post-gluconate treatments……………………………..…………………33
2.4 E. coli cell numbers pre-, during and post-gluconate treatmentss…………………...…………………………34
2.5 Percent Lactobacillus gluconate fermenters pre-, during and post-gluconate treatments ………………………………….…………...35
2.6 Percent Bifidbacterium gluconate fermenters pre-, during and post-gluconate treatments…………………………………………...…...36
2.7 Percent E. coli gluconate fermenters pre-, during and post-gluconate treatments……………………………………………….37
3.1 Alignment example of gluconate permease amino acid sequences from eight different bacteria using the DNAStar MegAlign software…..…………………………………43
3.2 Colony hybridization experimental design………………………….…...…47
3.3 Plasmid pORI28 ligated with putative gntK and gntP fragments transformed in E. coli EC1000………………………...54
3.4 Transformation of pTRK669 into L. reuteri 100-23……………………....….55 x Figure Page
3.5 Transformation of pORI28:gntP construct into L. reuteri 100-23 containing pTRK669…………………………………..….…56
3.6 Loss of pTRK669 and homologous recombination of pORI28:gntP with chromosomal gene copy when cells are upshifted to 48°C…………………………………………………….57
3.7 Southern hybridization analysis of L. reuteri 100-23 using the putative gntK or gntP PCR product as the probe……………………………………………………………….…….64
3.8 StuI digests of L reuteri 100-23 mutants A, B and D probed with putative gntP fragment and pORI28…………………..……..69
3.9 Northern analysis of L. reuteri wild-type and 100-23D……………….…….70
A.1 pH monitored over time for slurry and cheese combinations……….……83
A.2 Combination 2 bacterial cell counts…………………………………….……84
A.3 Combination 5 bacterial cell counts ………………………….……………...85
A.4 Combination 8 bacterial cell counts …………………………………..….….86
A.5 Combination 9 bacterial cell counts…………………………………..……...87
A.6 Combination 10 bacterial cell counts ……………………...……………..….88
A.7 Free amino acids over time for slurry and cheese combinations..…..….....89
A.8 Organic acid analysis of slurry and cheese combination 2………...……....90
A.9 Organic acid analysis of slurry and cheese combination 5………..….…....91
A.10 Organic acid analysis of slurry and cheese combination 8………..…...…..92
A.11 Organic acid analysis of slurry and cheese combination 9……………..….93
A.12 Organic acid analysis of slurry and cheese combination 10……..……..….94
xi CHAPTER 1
LITERATURE REVIEW
Consumers are becoming more health conscious. Today, the demand for functional foods that provide a health benefit beyond basic nutrition is on the rise. Research indicates that bacteria such as Lactobacillus and Bifidobacterium species are capable of eliciting health benefits such as reduced incidence of lactose intolerance and diarrhea, improved immune response and reduced cancer risk (20). These and other beneficial bacteria naturally inhabit the human intestinal tract. To thrive in the intestine, the environment must be rich with the nutrients that these bacteria require. A class of functional foods includes foods containing these beneficial bacteria, or probiotic bacteria. In addition to probiotic food supplements, foods may also be supplemented with prebiotic components.
A prebiotic is a non-viable food component that promotes the growth of a limited number of beneficial intestinal bacteria (72). The present studies address the characterization of gluconate metabolism genes in Lactobacillus and the effect of gluconate on human intestinal microflora.
1 1.1. Bacterial gluconate transport and catabolism
Escherichia coli
In E. coli, gluconate in the environment enters the bacterium via one of several gluconate transporters and is converted to 6-phosphogluconate by a gluconate-specific kinase. 6-phosphogluconate can then enter the Entner-
Doudoroff or pentose phosphate pathway (Figure 1.1). The Entner-Doudoroff pathway includes two enzymes: 6-phosphogluconate dehydratase, encoded by the edd gene and 2-keto-3-deoxy-6-phosphogluconate (KDPG) aldolase encoded by the eda gene (11, 12, 71). The enzymes in the Entner-Doudoroff pathway are induced by the presence of gluconate.
There are two systems responsible for gluconate metabolism in E. coli.
The primary system, GntI, includes the gntT, gntU and gntK genes which code
for high- and low-affinity gluconate transporters and a thermoresistant kinase,
respectively. These three genes along with edd and eda are negatively regulated
by the gntR gene product. The presence of gluconate prevents binding of the
GntR protein to its binding site (43). The gntRKU genes form an operon and are
subject to cyclic-AMP dependent catabolite repression. The edd and eda genes
are not.
2
Figure 1.1: Entner-Doudoroff pathway, adapted from Peekhaus and Conway (43)
The secondary gluconate system, GntII, contains a high-affinity transporter encoded by gntW and a thermosensitive gluconate kinase encoded by gntV. This system is primarily used for L-idonate catabolism, with gluconate as an intermediate (4, 43). A fourth gluconate transporter, encoded by gntP, was identified by Klemm et al (26). Unlike the other three transporters (gntT, gntU
3 and gntW), this transporter is not inducible by gluconate and is constitutively
expressed.
E. coli and many other enteric bacteria utilize the Entner-Doudoroff
pathway for metabolism of gluconate. It has been suggested that those organisms
capable of fermenting gluconate, but lacking 6-phosphogluconate dehydratase
(edd), utilize the pentose phosphate pathway to metabolize gluconate (43).
Bacillus subtilis
Bacillus subtilis differs from E. coli in that it only possesses one system for
gluconate metabolism. The Bacillus system contains a kinase (GntK), regulator
(GntR), permease (GntP) and a protein of unknown function (GntZ) (15). Fujita
and Miwa (14) discovered that the GntR protein represses transcription of the gnt
operon by binding to an operator site near the promoter. In addition, it was
shown that the presence of gluconate or glucono-δ-lactone prevent the regulatory
protein from binding to the operator.
Other bacteria
A few reports describe gluconate metabolism in bacteria of human or
food origin. London (33) indicates that many strains of Streptococcus faecalis and
Lactobacillus casei can metabolize gluconate. In these organisms, it has also been shown that synthesis of the 6-phosphogluconate dehydrogenase was repressed
4 by 20mM glucose (33). Bernsmann et al (5) showed that an inducible PEP-
dependent phosphotransferase specific for gluconate was present in Streptococcus
faecalis. London and Chace (34) suggest that a similar transferase exists in L.
casei. Tetaud et al (64) identified the 6-phosphogluconate dehydrogenase gene
from Lactococcus lactis using E. coli mutants. This enzyme converts 6-
phosphogluconate to ribose-5-phosphate, which can then be metabolized by the
pentose phosphate pathway. Analysis of the protein product revealed two
conserved arginine residues that are involved in substrate (6-phosphogluconate)
and NADP+ binding.
A gluconate manufacturer states in their promotional materials that
Bifidobacterium species are able to metabolize gluconate and that gluconate
consumption increases Bifidobacterium numbers in the intestine
(http://www.pmpinc.com/topics/topic2.html). No data or accessible citation
was provided along with this material; thus, it is difficult to assess the scientific validity of the claims.
1.2. Human intestinal microflora
Bacteria are natural inhabitants of the human gastrointestinal (GI) tract.
The bacterial GI population consists of a wide variety of bacterial species. The composition and quantity of bacteria vary from person to person and within the same person over time (19, 20, 57, 58, 72). Some bacterial species are transient
5 whereas others persist for long periods of time. Human bacterial populations can
reach 1010 colony forming units per gram of intestinal or fecal contents (43, 58).
Intestinal bacteria may be beneficial, neutral or harmful to the host. Purported benefits of consuming exogenous beneficial bacteria, or probiotic bacteria, include stimulation of the immune system, prevention of rotavirus diarrhea, reducing lactose intolerance and balancing intestinal microflora (20, 58).
In the early twentieth century, Metchnikoff first postulated the importance of lactobacilli in GI health (20). Intestinal inhabitants of this genus include
Lactobacillus acidophilus, L. brevis, L. casei, L. crispatus, L. fermentum, L. gasseri, L. johnsonii, L. reuteri, L. rhamnosus and L. salivarius (20, 57). In addition, intestinal
Bifidobacterium species are widely considered to be beneficial to their hosts (20).
Limited research on Propionibacterium species suggests that these bacteria may also inhabit the intestine and provide benefits (7, 44).
The factors affecting intestinal microflora composition in a given individual are poorly understood. Age, health status and diet may be involved, though substantial scientific evidence is lacking in these areas. Knowledge of how these organisms colonize and persist in the intestinal tract is minimal.
Adhesion to intestinal epithelial cells and competition between organisms for nutrients may play a role in which organism persists. Currently, researchers know little about what nutrients allow for intestinal colonization of a bacterium or what metabolic pathways and genes are important.
6 1.3. Role of E. coli gluconate metabolism and colonization of mouse intestine
Microorganisms in the gastrointestinal tract are extremely diverse. Little is known about how the bacteria compete with one another or what metabolic pathways or genes are important for colonization. E. coli persistence in the
human gastrointestinal tract varies by strain. Some strains will persist for years,
whereas others will persist for only days (51). Studies with E. coli indicated that
those strains capable of fermenting gluconate are better able to persist in the
mouse GI tract (55, 56). The mouse intestine has a gluconate concentration of 0.69
mM (43). E. coli F-18 was isolated in 1977 and shown to be an excellent colonizer
of the mouse intestine (42). Sweeney et al (55) found that the transporter GntP
allowed for excellent colonization of the F-18 strain in the mouse intestine.
Sweeney et al (55, 56) also showed that E. coli mutants of edd and eda will not
colonize the mouse large intestine. In addition, when the mutants were
complemented with plasmids containing functional edd and eda genes, the
colonization ability was restored. These findings strongly suggest that gluconate
metabolism confers an advantage for bacterial colonization of the gastrointestinal
tract.
7 1.4. The effect of dietary components on intestinal microflora.
Certain foods or food additives may have an impact on the diversity and
quantity of beneficial bacteria in the colon. If an intestinal bacterium can utilize a
particular unabsorbed food component better than its neighbors, that bacterium
may out-compete other flora and predominate (43, 55). Undigestible oligosaccharides, in particular fructooligosaccharides (FOS), have received
attention as prebiotic compounds. Foods such as artichokes, garlic and onion contain large quantities of FOS (16). FOS, as well as another carbohydrate inulin, have been shown to stimulate bifidobacterial growth within the intestine (16, 25,
46). Studies investigating the role of other potential prebiotic dietary components are lacking.
Dietary components may also have a negative impact on the overall intestinal microflora. Components that stimulate detrimental bacteria or that inhibit or kill beneficial bacteria may have a negative effect on the host. For example, fat source and quantity in the diet has been associated with detrimental changes in the intestine and inhibition of lactic acid bacteria. High fat diets with cocoa butter or beef fat alter bacterial enzymes in the rat intestine when compared to a low fat diet (38). Beef fat reduces bacterial numbers in the cecum compared to the low fat and cocoa butter diets. Fish fed different polyunsaturated fatty acids differ in viable lactic acid bacteria counts in the gut with linoleic acid reducing the counts most dramatically (45).
8 1.5. Gluconate as a potential prebiotic dietary component
Gluconate is generally recognized as safe (GRAS) by the FDA. It is a
natural component of some food (13) and is a common food ingredient.
Gluconate salts (Ca, Cu, Fe, K, Mg, and Zn) are added to foods or dietary
supplements as mineral sources due to their high solubility and stability.
Gluconate and glucono-δ-lactone (an ingredient that slowly releases gluconate)
are used as acidulants in dairy and meat products, sequestrants to limit lipid
oxidation, and flavoring (sour) agents. Calcium gluconate is added as the
calcium source in some calcium fortified foods and beverages. Gluconate is
likely to be present in the human large intestine from unabsorbed dietary
gluconate or from dead epithelial cells that contain 6-phospogluconate (55). A
gluconate manufacturer claims that only 20% of dietary gluconate is absorbed by
the small intestine, leaving 80% to proceed to the large intestine
(http://www.pmpinc.com/topics/topic2.html). Scientific validation of this
claim is necessary.
Published scientific studies, preliminary data in our laboratory and a gluconate manufacturer’s claims combine to suggest that this dietary component may impact beneficial intestinal bacteria. Based on the information available, it can be hypothesized that beneficial intestinal bacteria capable of fermenting gluconate may be better able to persist in the intestine and that dietary gluconate may promote the growth of indigenous gluconate-fermenting bacteria in the
9 intestine. Well-designed studies using human subjects are necessary to confirm
these suggestions.
1.6. Molecular biology approaches to assess complex bacterial communities
The human gastrointestinal tract contains a wide variety of
microorganisms. The intestinal microflora can achieve levels of 1010 colony
forming units per gram of intestinal or fecal contents (43, 58). The study of intestinal microorganisms relies on quantitative culturing of the organisms from feces. However, isolation from feces has a 50-80% recovery rate of the total population (60). Some microorganisms may not be culturable. Additionally,
numerous selective media must be employed to distinguish and quantitate
different microorganisms. This approach is cumbersome and may yield
unreliable information about the species and strains present. Therefore, new
molecular biology techniques such as pulsed-field gel electrophoresis (PFGE)
and denaturing gradient gel electrophoresis (DGGE) have been developed to
assess gastrointestinal microbial populations.
PFGE allows characterization of individual bacteria after selection on
selective medium. The method is based on rare-cutting restriction enzyme
digestions of genomic DNA (39). These enzymes digest genomic DNA into large
pieces that can be separated on an agarose gel. Unlike conventional gel
electrophoresis, PFGE utilizes alternating current to allow separation of the large
10 DNA fragments. Many studies have used PFGE to analyze changes in bifidobacterial and lactobacilli populations from human fecal samples (24, 40).
The disadvantage to PFGE is that it cannot monitor entire microbial populations, but rather individual bacteria that are first isolated on selective medium.
Unlike PFGE, DGGE can be used to monitor entire microbial populations without culturing or separating the organisms. DGGE separates DNA fragments of the same length, but containing different sequences based on differential denaturation characteristics of DNA fragments with different sequences. DNA is isolated directly from the fecal sample. A specific region of the DNA is amplified by polymerase chain reaction (PCR) and electrophoresed through a polyacrylamide gel with a linear gradient of DNA denaturant (41). Denaturants commonly used are mixtures of urea and formamide. Fragments of the same length with different sequences will denature into different configurations and migrate at different rates through the gel. Specific PCR primers are chosen or developed for investigation of specific populations. Or, less specific primers can be selected for a more global view of the population. Bands of interest can be excised from the gel for DNA sequencing and further study.
The PCR-DGGE method has been useful in assessing changes in microbial populations over time or during a treatment in samples that contain unculturable, difficult to culture or unknown bacteria. Samples such as water, soil, cheese, and feces have been investigated using this method. The method
11 has been applied to monitor or identify beneficial bacteria in human fecal samples. Satokari et al. (50) identified Bifidobacterium species from human feces by amplifying a 520 bp fragment from the 16S rRNA gene. Using primers,
Bif164-f and Bif662-GC-r, they were able to identify seven species of bifidobacteria from adult fecal samples. Walter et al (68) derived group specific primers that allow differentiation of Lactobacillus, Pediococcus, Leuconostoc and
Weissella species in human feces using the primer pair Lac1 and Lac2-GC, which amplify a 340 bp region of the 16Ss rRNA gene.
1.7. Rationale and Significance
The demand for functional foods, or those foods that elicit healthy benefits, is increasing. Consumers are more interested in how food components impact their health. Gluconate salts are common as an additive in several food products. If the beneficial bacterial populations in the intestine increase in numbers upon consumption of gluconate, consumers will have an alternative prebiotic food ingredient from which to choose, and ingredient suppliers and food manufacturers will have an alternative marketing strategy for their products. Knowledge of gluconate fermentation in Lactobacillus will aid researchers in their understanding of probiotic organisms and the natural human intestinal microflora.
12 1.8. Hypothesis and Goals of Project
Persistence of probiotic bacteria in the human gastrointestinal tract is influenced by numerous factors, including carbon source availability. Gluconate, a carbon source, is found in the gastrointestinal tract as well in food products.
Research studies have shown that genes involved in gluconate metabolism are needed for the persistence of E. coli in the mouse intestinal tract. This work is based on the hypothesis that gluconate aids in the persistence of beneficial bacteria in the human gastrointestinal tract. The specific goals of this work were to identify a gluconate-fermenting lactobacilli, characterize the gluconate genes within a chosen bacterial strain, generate a gluconate-negative mutant, evaluate enzyme activities in the wild-type and mutant strains and determine the effect of gluconate on the persistence of lactobacilli, bifidobacteria, propionibacteria and
E. coli in the human intestine.
13 CHAPTER 2
THE EFFECT OF DIETARY GLUCONATE ON FECAL CONTENT OF HUMAN LACTOBACILLUS, BIFIDOBACTERIUM, PROPIONIBACTERIUM AND ESCHERICHIA ISOLATES.
2.1. Abstract
Gluconate is naturally present in foods and is also used as a food additive and in dietary calcium supplements. The objective of this study was to assess the effect of dietary gluconate supplementation on selected bacterial populations from human feces. Twelve subjects consumed a daily calcium supplement. Calcium carbonate (control) was consumed for weeks 1 and 2, calcium gluconate (test) for weeks 3-5, and calcium carbonate for weeks 6 and 7. Weekly stool samples were collected, diluted and plated to enumerate Lactobacillus, Bifidobacterium,
Propionibacterium, and Escherichia coli. Colonies were subsequently grown on medium containing glucose or gluconate as the primary carbon source. At the level tested, dietary calcium gluconate did not significantly affect the overall quantity of Lactobacillus, Bifidobacterium, E. coli or Propionibacterium in most of the subjects tested. However, the proportion of gluconate-fermenting Lactobacillus
14 and Propionibacterium strains increased upon gluconate consumption in some subjects, suggesting that a shift in the species composition may occur. Pulsed
field gel analysis revealed 3-8 distinct banding patterns for each subject’s gluconate-fermenting lactobacilli. Speciation using the 16S-23S rRNA intergenic spacer region showed 2-6 different gluconate-fermenting Lactobacillus species in
each subject and 11 different species overall. The majority of the gluconate-
fermenting lactobacilli were L. rhamnosus or L. paracasei.
2.2. Introduction
Microorganisms in the gastrointestinal tract are extremely diverse. Little is
known about how the bacteria compete with one another or what metabolic
pathways or genes are important for colonization and persistence. Escherichia coli persistence in the human gastrointestinal tract varies by strain. Some strains persist for years, whereas others will only remain for days (51). Studies with E.
coli indicate that those strains capable of fermenting gluconate are better able to
persist in the mouse gastrointestinal tract (55, 56). The mouse intestine has a
gluconate concentration of 0.69 mM (43). Escherichia coli F-18 was isolated in 1977
and shown to colonize the mouse intestine efficiently (42). Sweeney et al (55)
found that the gluconate transporter GntP facilitated this colonization and that
edd and eda mutants did not colonize the mouse large intestine (55, 56). The edd and eda loci encode the 6-phosphogluconate dehydratase and 2-keto-3-deoxy-6- phophogluconate aldolase, respectively. In addition, when the mutants were 15 complemented with plasmids containing functional edd and eda genes,
colonization was restored. These findings strongly suggest that gluconate
metabolism confers an advantage for E. coli colonization of the gastrointestinal
tract.
Certain foods or food additives may have an impact on the diversity and
quantity of beneficial bacteria in the colon. If an intestinal bacterium can utilize a
particular unabsorbed food component better than its neighbors, that bacterium
may out-compete other flora and predominate (43, 55). Undigestible oligosaccharides, in particular fructooligosaccharides (FOS), have received
attention as prebiotic compounds. Studies investigating the role of other
potential prebiotic dietary components are limited. Additionally, dietary
components may also have a negative impact on the overall intestinal microflora
and on the host organism. These components may stimulate detrimental bacteria
or inhibit or kill beneficial bacteria. Gluconate is generally recognized as safe
(GRAS) by the FDA. It is a natural component of some foods (13) and is a
common food ingredient. Gluconate salts (Ca, Cu, Fe, K, Mg, and Zn) are added
to foods or dietary supplements as mineral sources due to their high solubility
and stability. Gluconate and glucono-δ-lactone (an ingredient that slowly
releases gluconate) are used as acidulants in dairy and meat products,
sequestrants to limit lipid oxidation, and flavoring agents. Gluconate is likely to
be present in the human large intestine from unabsorbed dietary gluconate or
16 from dead epithelial cells that contain 6-phospogluconate (55). Published studies
(2, 33) and data from our laboratory (see Chapter 3) suggest that this dietary component may support the growth of beneficial intestinal bacteria. Therefore, it is the purpose of this study to evaluate the gluconate-fermenting lactobacilli, bifidobacteria, propionibacteria and E. coli populations of the human gastrointestinal tract before, during and after oral gluconate supplementation, and to characterize gluconate-fermenting human Lactobacillus isolates.
2.3. Materials and Methods
Subject selection, treatment and study design
Seventeen healthy, non-smoking human subjects were recruited from The
Ohio State University (OSU) campus and the city of Columbus, Ohio. Approval for human subjects testing was acquired under OSU protocol number
2003H0088. Subjects were screened to eliminate users of antibiotics and those with recurrent intestinal complaints. For the duration of the study, subjects were instructed to avoid consumption of foods and supplements containing added gluconate salts or glucono-δ-lactone other than what is administered by the research team. Subjects were also asked to avoid foods and supplements containing probiotic bacteria (such as yogurt). Commercially available calcium supplements containing calcium gluconate (Freeda vitamins, New York, NY) were used as the dietary gluconate source. The dosage delivered 2.4 grams of 17 gluconate per day. To control for any effect of calcium supplementation, subjects
consumed supplements containing an equivalent mass of calcium in the form of
calcium carbonate during the pre- and post-gluconate treatment periods. The
experiment was designed as a crossover study. Therefore, stool samples were
collected from each subject once per week prior to gluconate supplementation
(two weeks), during gluconate supplementation (three weeks) and after gluconate supplementation (two weeks). The timeline of collection is illustrated
in Figure 2.1. Sample collection containers were provided (catalog #02-544-208,
Fisher Scientific, Pittsburgh, PA), and subjects were asked to refrigerate samples
immediately after collection and to deposit labeled samples in a designated
laboratory refrigerator within 1 hour after collection.
Calcium Carbonate Calcium Gluconate Calcium Carbonate
Supplement Supplement Supplement 14 days 21 days 14 days
Stool sample collection and analysis
Figure 2.1: Calcium supplementation and sampling time line.
18 Quantitation of bacterial population
All fecal samples were placed in an Forma Scientific anaerobic chamber (85% nitrogen, 10% hydrogen, 5% carbon dioxide) and processed anaerobically immediately upon receipt. All reagents and media were pre-reduced by placing in the anaerobic chamber overnight. One gram of sample was homogenized in pre-reduced 0.1% peptone and serially diluted in the same medium. Diluted samples were plated on selective media and incubated for the indicated durations and temperatures (Table 2.1). After incubation, colony-forming units per gram (cfu/g) were counted for each type of bacterium. To determine the percentage of each group able to ferment gluconate, 50 to 100 colonies were selected randomly at each time point and streaked with a sterile toothpick onto medium containing glucose (100 mM) and medium containing gluconate (100 mM) as the sole carbon source (see Table 2.1 for specific media). Non-gluconate fermenters were able to grow on MRS and modified sodium lactate agar containing gluconate, therefore bromcresol purple (BCP) was added to these agars as a pH indicator. Those capable of fermenting gluconate turned yellow, whereas those incapable of fermenting gluconate remained white. Results on each medium were verified with selected strains on chemically defined medium.
The percentage of gluconate fermenters was calculated as the number of colonies that grew on the gluconate medium divided by the number of colonies that grew on the glucose medium multiplied by 100.
19
Bacterial Group Medium Incubation Medium for gluconate-fermenting Analyzed for total duration percentage counts and conditions E. coli Brilliant 1 day, 37°C, M63 M63 (glucose) (31) Green aerobic (gluconate) Bile agara Lactobacillus Rogosa 2 days, Lactobacilli Lactobacilli (59) SL agarb 37°C, MRSb BCPc agar MRS BCP agar anaerobic (gluconate) (glucose) Bifidobacterium Rogosa 4 days (after Lactobacilli Lactobacilli (52,59) SL agar Lactobacillus MRS BCP MRS BCP colonies are 0.05% cysteine- 0.05% cysteine- marked at 2 hydrochloride hydrochloride days), 37°C, agar (gluconate) agar (glucose) anaerobic Propionibacterium Lithium 6 days, Modified Modified (10,37) Glycerol 30°C, sodium lactate sodium lactate agar anaerobic agard agar (glucose) (gluconate) a Alpha BioSciences, Baltimore, MD b Difco, Becton, Dickinson and Company, Franklin Lakes, NJ c Bromcresol purple, pH indicator d Sodium gluconate or glucose was used in place of sodium lactate
Table 2.1: Media and growth conditions used to enumerate bacterial cell counts and percent gluconate fermenters in fecal samples
20 Molecular typing and speciation of gluconate-fermenting Lactobacillus species
Thirty gluconate-fermenting Lactobacillus isolates from each subject and one non-gluconate-fermenting isolate from nine subjects were analyzed by pulsed field gel electrophoresis (PFGE) as described by Jenkins et al. (23). Briefly,
DNA from Lactobacillus isolates was prepared, digested with ApaI and electrophoresed. PFGE conditions were as follows: 1% PFGE agarose gel, 0.5 x
TBE buffer, 5 V/ cm-1 and 1-12 seconds switching time for 14 hours, followed by
0.5-2 seconds switching time for 3 hours. DNA fragments were visualized by ethidium bromide staining followed by UV transillumination.
Three to thirteen distinct banding patterns were observed in all subjects.
Each of these isolates was speciated by amplifying the spacer region between the
16S and 23S rRNA genes using the method of Tannock et al. (63) Briefly, chromosomal DNA from specified isolates was subjected to PCR using the
Expand High Fidelity PCR System (Roche Applied Science, Indianapolis, IN) and the following primer set: 16-1A (5’-GAATCGCTAGTAATCG-3’) and 16-1B (5’-
GGGTTCCCCCATTCGGA-3’). PCR products were electrophoresed through a
1% agarose gel, stained in ethidium bromide solution and visualized by UV transillumination. The band located at 500-600 bp was excised and purified using the Qiaquick gel extraction kit (Qiagen, Valencia, CA). Purified fragments were sequenced using primer 16-1A (63) and BigDye Terminator Cycle
Sequencing chemistry in a 3700 DNA analyzer (Applied Biosystems, Foster City,
21 CA) in the Plant-Microbe Genomics Center at The Ohio State University
(Columbus, OH). DNA sequences were compared to those available in the
GenBank database on October 19, 2004 using the BLASTN algorithm (1).
Sequences of greater than or equal to 97.5% similarity were used to determine
species identity of the isolate.
Statistical analysis
Results from each treatment (pre-, during and post-gluconate
supplementation) were averaged and compared within each subject using one-
way ANOVA and Tukey’s post-hoc test at the 95% confidence level (SigmaStat
version 3.1, Systat Software, Inc., Richmond, CA). Values with P<0.05 were considered statistically different.
2.4. Results
Human microflora: Five of the 17 subjects were disqualified or did not complete the study. The remaining 12 completed the entire study adhering to all guidelines. The data collected demonstrated that at the level tested, dietary gluconate supplementation had no significant effect on the overall population of beneficial bacteria or E. coli in most subjects. All 12 subjects had lactobacilli in their stool samples. Total Lactobacillus counts did not change significantly throughout the study in all but two subjects (Figure 2.2). In subject 998,
22 Lactobacillus numbers decreased upon gluconate supplementation; whereas, subject 059 showed a significant increase in lactobacilli during gluconate supplementation. Ten of the subjects had bifidobacteria during the study (Figure
2.3). Subject 059 also showed a significant increase in Bifidobacterium counts upon gluconate supplementation, whereas others showed no significant change.
Eleven of the 12 subjects had E. coli throughout the study (Figure 2.4). Escherichia coli counts decreased in subject 478 upon gluconate supplementation, and no significant change was observed in the other subjects. Eleven of the 12 subjects had propionibacteria sporadically throughout the study (data not shown).
Gluconate-fermenting lactobacilli were isolated from all subjects (Figure
2.5). The percentage of lactobacilli able to ferment gluconate ranged from 1.8 to
100% over all time points and subjects. Percentages increased significantly during gluconate consumption in subjects 059 and 382 and remained unchanged in the other 10 subjects. Gluconate-fermenting bifidobacteria were recovered from 7 subjects with 1-100% able to ferment gluconate (Figure 2.6). Subject 250 had significantly reduced gluconate-fermenting bifidobacteria during gluconate treatment. Most E. coli isolates were able to ferment gluconate (Figure 2.7). There was no significant change in the percentage of E. coli able to ferment gluconate during gluconate supplementation. In 8 subjects, gluconate-fermenting propionibacteria were found at levels of 38 – 100% of the population (Table 2.2).
23 Gluconate-fermenting propionibacteria were only encountered after gluconate
consumption began in 6 of these subjects.
Subject Pre-gluconate Percent During-gluconate Post-gluconate Fermenters Percent Fermenters Percent Fermenters 059 NDa ND ND 243 81% 100% 100% 250 100% 100% 100% 305 ND ND 100% 382 ND 38% ND 433 ND 50% ND 478 ND ND ND 494 ND ND ND 587 ND ND ND 881 ND 100% 100% 901 ND 92% ND 998 ND 100% 100% a No gluconate fermenters detected
Table 2.2: Percent gluconate-fermenting propionibacteria pre-, during and post- gluconate treatments
Characterization of lactobacilli: Gluconate-fermenting and non-fermenting (1
each from 9 subjects) Lactobacillus colonies from each subject at various time-
points were subjected to pulsed field gel electrophoresis (PFGE) analysis to
separate genetically different isolates prior to speciation and evaluate the
diversity of the population. A total of 87 different banding patterns (gluconate-
24 fermenting isolates) from all subjects were noted by PFGE analysis. These 87
banding patterns were speciated using the intergenic region of the 16S-23S rRNA
gene. From the speciation, a total of 11 different gluconate-fermenting species
were found in all subjects (Table 2.3). Most subjects’ gluconate-fermenting
lactobacilli were of the species rhamnosus or paracasei.
2.5. Discussion
There are few studies reporting the effect of gluconate on intestinal bacterial cell number, and no studies reporting the number of gluconate- fermenting bacteria in the human gastrointestinal tract.
This study illustrated subject-dependent population changes in bacterial numbers upon gluconate supplementation. Most cell numbers did not significantly change during gluconate supplementation. Asano et al. (2) found that administration of glucono-δ-lactone significantly increased the number of bifidobacteria in 10 subjects, had no effect on the number of lactobacilli and decreased the number of Clostridium perfringens. The exact gluconate dosage in the Asano et al study is not known because these authors administered glucono-
δ-lactone (9 and 3 g/day) in contrast to the present study where a gluconate salt
(2.4 g of gluconate/day) was administered.
Variation in bacterial populations in human feces between and within subjects has been demonstrated in numerous studies. For example, Kimura et
25 al. (24) examined the fecal populations of bifidobacteria and lactobacilli with no
dietary modulation except the absence of yogurts and antimicrobial therapies. As
in the present study, lactobacilli cell numbers varied greatly between subjects
and also between fecal samples within the same subject (104-108 cfu/g). Genetic analysis revealed that 8 of the 9 subjects in which lactobacilli were detected had four or less species.
Previous studies have demonstrated the effect of administering oral prebiotic carbohydrates and probiotic bacteria. Tannock et al. (61) demonstrated the effect of oral consumption of biscuits containing galacto- or fructo- oligosaccharides on faecal microflora using nucleic acid based methods. The authors found no significant increase in bifidobacteria cell numbers, but an increase in their metabolic activity upon consumption of the biscuits. In contrast,
Tuohy et al (67) found an increase in bifidobacteria cell numbers during oral administration of partially hydrolyzed guar gum and fructo-oligosaccharides.
Bougle et al. (7) found that orally administered propionibacteria do survive the
human stomach, but do not colonize in gastrointestinal tract. These authors also
found that some propionibacteria species promote the growth of bifidobacteria
species in the intestine. Huang et al. (21) found only one Propionibacterium strain
out of 13 that adheres to human intestinal epithelial cells in vitro. These studies
support the present data of sporadic propionibacteria cell numbers.
26 To our knowledge, there are no reports of gluconate-fermenting bacteria
isolated from humans. The number of isolates able to ferment gluconate
increased in 2 and 6 subjects for lactobacilli and propionibacteria bacterial
populations, respectively, upon consumption of the gluconate supplement. This
suggests that although overall bacterial cell numbers did not increase
significantly, the distribution within these genera may change due to dietary
gluconate. The present study represents a comprehensive survey of gluconate
fermentation in human lactobacilli. Most gluconate-fermenting isolates found in
this study were of the species paracasei or rhamnosus. In addition, like Asano et al.
(2), a few isolates of gluconate-fermenting L. fermentum and L. casei were found in the present study.
If gluconate does provide an ecological niche for beneficial bacteria, they might be better able to persist in the human gastrointestinal tract with dietary gluconate supplementation. A gluconate-fermenting probiotic strain that is able to utilize gluconate at a more rapid rate than that of non-beneficial microbes may be able to out compete other intestinal inhabitants to become better established.
With this information, food manufacturers could add gluconate to food products as a dietary source or provide gluconate in those foods that already contain probiotic organisms.
A more thorough understanding of gastrointestinal microflora interactions is needed. Studies of in vitro human epithelial cells are beneficial, however a real
27 representation of the human gastrointestinal tract in which researchers could manipulate conditions and microbial interactions would be useful in determining the power of probiotic strains and prebiotic compounds.
28
Isolate code Non-gluconate fermenters 243 Lactobacillus rhamnosus 382 Pediococcus acidilactici 433 Lactobacillus acidophilus 478 Lactobacillus paracasei subsp. paracasei 494 Lactobacillus paracasei subsp. paracasei 682 Lactobacillus. fermentum 881 Lactobacillus paracasei subsp. paracasei 901 Enterococcus faecium 998 Enterococcus faecalis Gluconate fermenters 59 3Q Enterococcus faecium 59 4A Lactobacillus paracasei subsp. paracasei 59 4B Lactobacillus rhamnosus 59 4C Lactobacillus paracasei subsp. paracasei 59 4F Lactobacillus paracasei subsp. paracasei 59 5A Lactobacillus paracasei subsp. paracasei 59 5B Lactobacillus fermentum 250 0B Lactobacillus rhamnosus 250 2C Lactobacillus paracasei subsp. paracasei 250 3L Lactobacillus fermentum 250 4A Lactobacillus rhamnosus 250 6K Lactobacillus rhamnosus 305 4A Lactobacillus paracasei subsp. paracasei 305 4D Lactobacillus paracasei subsp. paracasei 305 5A Lactobacillus rhamnosus 305 5B Lactobacillus paracasei subsp. paracasei 305 7A Weisella kimchii 305 7C Lactobacillus rhamnosus 382 2D Lactobacillus rhamnosus 382 2J Lactobacillus rhamnosus 382 3J Lactobacillus rhamnosus
Continued
Table 2.3: Lactobacillus isolates determined by sequencing the 16S-23S rRNA intergenic region
29 Table 2.3 continued
382 3T Lactobacillus fermentum 382 5A Lactobacillus paracasei subsp. paracasei 382 5N Lactobacillus rhamnosus 382 6A Lactobacillus fermentum 382 6H Lactobacillus paracasei subsp. paracasei 433 6A Lactobacillus rhamnosus 433 7A Lactobacillus rhamnosus 433 7B Lactobacillus fermentum 478 0A Lactobacillus rhamnosus 478 1J Lactobacillus hilgardii 478 1K Lactobacillus rhamnosus 478 1P Lactobacillus rhamnosus 478 3F Lactobacillus paracasei subsp. paracasei 478 4A Enterococcus dispar 478 4R Lactobacillus rhamnosus 478 5N Lactobacillus paracasei subsp. paracasei 478 6A Lactobacillus rhamnosus 478 6B Lactobacillus rhamnosus 478 7J Enterococcus faecalis 478 7P Lactobacillus rhamnosus 494 0E Lactobacillus paracasei subsp. paracasei 494 7F Lactobacillus paracasei subsp. paracasei 494 7N Lactobacillus paracasei subsp. paracasei 494 7R Lactobacillus paracasei subsp. paracasei 494 7T Lactobacillus paracasei subsp. paracasei 587 0K Lactobacillus rhamnosus 587 4B Lactobacillus rhamnosus 587 4S Lactobacillus rhamnosus 587 6E Lactobacillus rhamnosus 587 6P Lactobacillus rhamnosus 682 0B Lactobacillus rhamnosus 682 1T Lactobacillus paracasei subsp. paracasei 682 2A Lactobacillus paracasei subsp. paracasei 682 3L Lactobacillus rhamnosus
Continued
30 Table 2.3 continued
682 7A Lactobacillus fermentum 682 7K Lactobacillus rhamnosus 881 7B Lactobacillus rhamnosus 881 6O Lactobacillus rhamnosus 881 6N Lactobacillus plantarum 881 6F Lactobacillus paracasei subsp. paracasei 881 5A Lactobacillus rhamnosus 881 4L Lactobacillus rhamnosus 881 4J Lactobacillus rhamnosus 881 3B Lactobacillus paracasei subsp. paracasei 881 2E Lactobacillus paracasei subsp. paracasei 881 2D Lactobacillus paracasei subsp. paracasei 881 2A Lactobacillus rhamnosus 881 0G Lactobacillus plantarum 881 0F Lactobacillus rhamnosus 901 1A Lactobacillus rhamnosus 901 2A Lactobacillus rhamnosus 901 3A Lactobacillus paracasei subsp. paracasei 901 3L Lactobacillus paracasei subsp. paracasei 901 7A Lactobacillus rhamnosus 901 7B Lactobacillus rhamnosus 998 1A Lactobacillus rhamnosus 998 1C Lactobacillus rhamnosus 998 2A Weisella kimchii 998 2B Weisella confusa 998 2E Weisella confusa 998 2F Weisella confusa 998 4A Lactobacillus rhamnosus 998 6C Lactobacillus rhamnosus 998 6E Lactobacillus rhamnosus
31
9
8 059 250 7 305 382 6 433 478 5 494 log cfu/g log 587 4 682 881 3 901 998 2
pre during post Treatment
Figure 2.2: Lactobacillus cell numbers pre-, during and post-gluconate treatments.
32
10
9
8
7 059 250 6 305 382 log cfu/g 5 433 478 494 4 682 881 3 901
2 pre during post
Treatment
Figure 2.3: Bifidobacterium cell numbers pre-, during and post-gluconate treatments.
33
9
8
7
059 6 250 305 382
log cfu/g log 5 433 478 4 587 682 881 3 901 998 2 pre during post Treatment
Figure 2.4: E. coli cell numbers pre-, during and post-gluconate treatments.
34
pre 100 during post 80
60
40
Fermenters Gluconate Percent 20
0 059 250 305 382 433 478 494 587 682 881 901 998
Figure 2.5: Percent Lactobacillus gluconate fermenters pre-, during and post- gluconate treatments.
35
100 pre during post 80
60
40
Percent Gluconate Fermenters 20
0 059 250 305 382 433 478 494 682 881 901
Figure 2.6: Percent Bifidobacterium gluconate fermenters pre-, during and post- gluconate treatments.
36
100
80
60
40
Percent Gluconate Fermenters 20
0 059 250 305 382 433 478 587 682 881 901 998 pre during post
Figure 2.7: Percent E. coli gluconate fermenters pre-, during and post-gluconate treatments.
37 CHAPTER 3
GENETIC CHARACTERIZATION OF GLUCONATE GENES IN LACTOBACILLUS REUTERI 100-23
3.1. Abstract
The objective of this study was to characterize gluconate metabolism
genes in Lactobacillus reuteri 100-23. A 500 base pair (bp) putative gluconate
kinase fragment and a 750 bp putative gluconate permease fragment were found
by polymerase chain reaction using degenerate primers. Subsequent colony
hybridization experiments were unsuccessful; therefore complete gluconate
metabolism gene sequences in L. reuteri 100-23 could not be elucidated. A
gluconate negative mutant of L. reuteri 100-23 was constructed via homologous
recombination utilizing the putative gluconate permease fragment. The mutant
(100-23D) and wild-type strains were characterized for gluconokinase, 6-
phosphogluconate dehydrogenase and gluconate uptake activities in MRS and
chemically defined media (CDM). Gluconokinase activities of the wild-type were
significantly higher in cultures grown in CDM-gluconate than in MRS-gluconate.
Lactobacillus reuteri 100-23D had significantly lower gluconokinase and gluconate
38 uptake activities compared to wild-type in both media. There was no significant
difference in 6-phosphogluconate dehydrogenase activity between wild-type and
100-23D in MRS broth. Gluconate uptake and gluconokinase enzyme analysis
indicated that gluconate metabolism genes are induced by the presence of
gluconate and possibly arranged in an operon. Transcriptional analysis of wild-
type and mutant strains showed a single transcript. Transcriptional analysis also
indicated that transcription of the gluconate permease gene is not inducible, but
rather constitutively expressed in the presence of glucose or gluconate.
3.2. Introduction
Gluconate is a naturally occurring carbohydrate. Sources of gluconate in
the large intestine include foods, dietary supplements and dead epithelial cells.
Sweeney et al. (55, 56) demonstrated that E. coli mutants unable to utilize
gluconate were not able to colonize the mouse intestine, indicating that gluconate
utilization genes may be important in colonization by intestinal bacteria. The
gluconate genes of E. coli and B. subtilis have been extensively studied.
In E. coli, gluconate in the environment enters the bacterium via one of several gluconate transporters and is converted to 6-phosphogluconate by a gluconate-specific kinase. 6-phosphogluconate can then enter the Entner-
Doudoroff or pentose phosphate pathway. The Entner-Doudoroff pathway includes two enzymes: 6-phosphogluconate dehydratase, encoded by the edd
39 gene and 2-keto-3-deoxy-6-phosphogluconate (KDPG) aldolase encoded by the
eda gene (11, 12, 71).
In E. coli, there are two systems responsible for gluconate metabolism.
The primary system, GntI, includes the gntT, gntU and gntK genes which code
for high- and low-affinity gluconate transporters and a thermoresistant kinase,
respectively (22). These three genes along with edd and eda are negatively
regulated by the gntR gene product. The presence of gluconate prevents binding
of the GntR protein to its binding site (43). The gntRKU genes form an operon
and are subject to cyclic-AMP dependent catabolite repression. The edd and eda
genes are not arranged in an operon and are not sensitive to cyclic-AMP
dependent catabolite repression.
The secondary gluconate system, GntII, contains a high-affinity
transporter encoded by gntW and a thermosensitive gluconate kinase encoded by
gntV. This system is primarily used for L-idonate catabolism, with gluconate as
an intermediate (4, 43). A fourth gluconate transporter, encoded by gntP, was
identified by Klemm et al (26). Unlike the other three transporters (gntT, gntU
and gntW), gntP is not inducible by gluconate and is constitutively expressed.
Bacillus subtilis differs from E. coli in that it only possesses one system for
gluconate metabolism. The Bacillus system contains a kinase (GntK), regulator
(GntR), permease (GntP) and a protein of unknown function (GntZ) (15). Fujita and Miwa (14) discovered that the GntR protein represses transcription of the gnt
40 operon by binding to an operator site near the promoter. In addition, it was shown that the presence of gluconate or glucono-δ-lactone prevent the regulatory protein from binding to the operator.
Reports describing gluconate metabolism in bacteria of human or food origin are limited. London (33) indicated that many strains of Streptococcus faecalis and Lactobacillus casei can metabolize gluconate. The present study investigates gluconate utilization in L. reuteri.
3.3. Methods
Screening of bacteria for gluconate utilization
Seven Lactobacillus, twelve Propionibacterium and three Bifidobacterium strains from human and dairy origins were screened for their ability to use gluconate as a sole carbon source (see Table 3.2 for source of bacteria).
Lactobacillus strains were tested using MRS medium (Difco Lactobacilus MRS
Broth, Becton, Dickinson and Company, Franklin Lakes, NJ) in which the glucose was replaced with gluconate (20 g/L). Sodium acetate as well as residual glucose from medium components such as yeast and beef extracts allowed some growth of gluconate-negative colonies; however, gluconate-positive bacteria were easily differentiated by incorporating the pH indicator, bromcresol purple
(0.005%) into the medium. These results were confirmed in chemically defined medium (CDM) containing 20 g/L of gluconate as the sole carbon source (29).
41 Propionibacterium strains were tested in defined medium containing gluconate (12
g/L) instead of glucose (17). Bifidobacterium strains were tested in reinforced
clostridial medium (Becton, Dickinson and Company, Franklin Lakes, NJ) with
gluconate (5 g/L) replacing glucose. Lactobacillus and Bifidobacterium strains were
grown at 37°C, whereas Propionibacterium strains were grown at 30°C. All strains
were grown anaerobically (85% nitrogen, 10% hydrogen, 5% carbon dioxide) in a
Forma Scientific anaerobe system (model 1025). Cell growth was determined by
measuring the A600 of each sample in a Spectronic 20 spectrophotometer
(Spectronic Instruments Inc., Rochester, NY).
Primer Design and PCR conditions
Attempts at utilizing primer sets based on the gluconate genes of
Escherichia coli and Lactococcus lactis were unsuccessful in amplifying the
gluconate metabolism genes of Lactobacillus reuteri 100-23. Therefore, degenerate
primers were designed based on the alignment of the amino acid sequences of the gluconate kinase (gntK) gene from Bacillus subtilis (15), B anthracis,
Clostridium perfringens, L. lactis (6) and Listeria innocua. Degenerate primers were
also designed for the gluconate permease (gntP) gene using the amino acid
sequences from B. subtilis (15), B. licheniforms, E. coli (26), L. plantarum,
Staphylococcus aureus, Pseudomonas aeruginosa, Salmonella enterica and S.
42 typhimurium (see Figure 3.1 for an example alignment). The MegAlign program from DNAStar (Lasergene, Madison, WI) was used to find the best possible alignment. Table 3.1 lists degenerate primer sets utilized.
Genomic L. reuteri 100-23 DNA (2 µg) was subjected to PCR cycles using the following conditions: 94°C, 45 seconds; 45°C, 45 seconds; 72°C, 90 seconds for
40 cycles followed by 72°C for 10 minutes.
Figure 3.1: Alignment example of gluconate permease amino acid sequences from eight different bacteria using the DNAStar MegAlign software.
43 Primer Set Upstream Primer Downstream Primer (amino acid sequence) (amino acid sequence) Gluconate Kinase 1 ITWAD GTSGA 2 ITWAD FHPYL 3 ITWAD GERAP 4 GTSGA FHPYL 5 GTSGA GERAP Gluconate Permease 6 HGFLPPH IIGGGG 7 HGFLPPH GSATVA 8 HGFLPPH HVNDAGFW 9 HGFLPPH SHVND 10 IIGGGG GSATVA 11 IIGGGG HVNDAGFW 12 IIGGGG SHVND 13 GSATVA HVNDAGFW 14 GSATVA SHVND
Table 3.1: Amino acid sequences of degenerate primer sets used to search for L. reuteri 100-23 gluconate genes.
PCR products were cloned into plasmid pGEM-T (Promega, Madison, WI)
and sequenced using BigDye Terminator Cycle Sequencing chemistry and a 3700
DNA analyzer (Applied Biosystems, Foster City, CA) in the Plant-Microbe
44 Genomics Center at The Ohio State University (Columbus, OH). The amino acid
sequence, deduced from the DNA sequence, was compared to the GenBank
database using the BLASTX algorithm (1)
Southern analysis
To obtain entire gene sequences, L. reuteri 100-23 genomic DNA was
digested with EcoRI and four identical samples were electrophoresed on a 0.8%
agarose gel (SeaKem GTG Agarose, Cambrex Bio Science, Rockland, ME) along
with a molecular weight marker (1 Kb Plus DNA Ladder, Invitrogen, Carlsbad,
CA) and putative gntK and gntP fragments as controls (obtained from degenerate
primer experiments). The gel was cut and three of the four lanes containing the
100-23 genomic DNA were separated from the rest of the gel, wrapped in plastic
and stored at 4°C while the remaining portion of the gel was transferred to a
nylon membrane (MagnaCharge, Osmonics, Inc., Minnetonka, MN) by capillary
transfer (49). The putative gntK or gntP PCR product was labeled using the DIG
High Prime Labeling and Detection Starter Kit II (Roche Applied Sciences,
Indianapolis, IN). The membrane hybridization at 42°C (over night) and label detection were performed according to the kit manufacturer instructions.
Regions of similarity (2-4 kb) were excised from the refrigerated gel and gel-purified using the Qiaquick gel extraction kit (Qiagen, Valencia, CA). The products were then ligated into EcoRI-cut plasmid pBlueScript-KS+ to construct a
45 subgenomic library of L. reuteri 100-23. The ligation mix was then transformed
into E. coli DH5-α via electroporation (49). Transformants were selected on LB ampicillin (100 µg/ml) agar plates. Transformants with plasmid containing inserts were differentiated using IPTG and X-gal (49) spread on the LB ampicillin agar plates.
Colony hybridization
Individual colonies from the transformation were spotted onto duplicate
LB (49) ampicillin (100 µg/ml) agar plates and incubated for 18 h at 37°C, with
one plate being refrigerated (Figure 3.2, plate B) and the other used for colony
hybridization (Figure 3.2, plate A) using the putative gntP fragment as the
probe.
Colonies that were transformed with pBlueScript-KS+ sub-genomic L.
reuteri 100-23 constructs (Figure 3.2, plate A) were transferred to a nylon
membrane cut to fit within the Petri plate (MagnaCharge 82 mm; Osmonics, Inc.,
Minnetonka, MN). Membranes were carefully removed with forceps from the
pertri plate and placed colony side up on saturated 3M chromatography paper in
the following order: 10% SDS for 3 minutes; denaturing solution (0.5 N NaOH;
1.5 M NaCl) for 10 minutes, neutralization solution (1.5 M NaCl; 0.5 M Tris-Cl,
pH 7.4) for 10 minutes and 2× SSC for 10 minutes.
46 Membranes were then air-dried and cross-linked in an UV Stratalinker
1800 (Stratagene, La Jolla, CA) and then hybridized with digoxigenin-labeled putative gntP PCR product at 42°C over night. After hybridization, membranes
were washed at room temperature twice for 5 minutes with 2× SSC 0.1% SDS and
twice with 0.5× SSC 0.1% SDS for 15 minutes. Membranes were then subjected to
reagents in the DIG High Prime Labeling and Detection Starter Kit II (Roche
Applied Sciences, Indianapolis, IN)...... A B
Prepare membrane ... and hybridize to A . .. labeled probe ...... Detect labeled probe .
Figure 3.2 Colony hybridization experimental design. Plate “B” was placed at 4°C for later use while plate “A” was used for colony hybridization experiments. White circles denote nylon membrane and gray circles denote LB agar plates.
47 Enzyme Assays
Cell-free extracts were prepared by growing E. coli K12, wild-type L.
reuteri 100-23 and mutant L. reuteri 100-23D to mid-log phase in MRS or
chemically defined (CDM) medium containing gluconate (100 mM), glucose (100
mM) or gluconate + glucose (50 mM gluconate and 50 mM glucose) (29). Cells
were centrifuged and washed in breaking buffer (0.2 M Tris-HCl, pH 7.6, 0.2 M
KCl, 10 mM magnesium acetate, 0.3 mM DTT, 5% glycerol; (69). After washing,
the cells were resupsended in 1 ml of breaking buffer. A small amount of 0.1 mm
glass beads (BioSpec Products, INC, Bartlesville, OK) was added. Cells were
pulverized in a Mini Bead-Beater-8 (BioSpec Products, INC, Bartlesville, OK) for
30 seconds, 6 times with incubation on ice for 1 minute between each session to prevent excessive heating.
Protein concentration was measured in each cell-free extract by the
method of Bradford (8). Bio-rad dye reagent (Bio-Rad Laboratories, Hercules,
CA) was diluted 1:4 in water and filtered through Whatman #1 filter paper. Ten
µl of the cell-free extract were added to 200 µl of diluted dye reagent in a 96-well
microtiter plate. The plate was mixed on an orbital shaker for 5 minutes at room
temperature. The absorbance was then read at 490 nm in a microtiter plate reader
(Molecular Devices, Sunnyvale, CA). Bovine serum albumin (BSA) standards
were subjected to the same treatment. A standard curve was constructed from
48 the BSA concentrations and used to calculate the protein concentration of each
cell-free extract.
6-phosphogluconic acid dehydrogenase assay: The following reagents
were added to make a total volume of 1.0 ml and a final concentration of 50 mM
HEPES, pH 7.6, 10 mM MgCl2, 500 µM 6-phosphogluconic acid, 0.2 mM NADP
and 10 µl cell-free extract. Reactions were started by the addition of the cell-free
extract. All reactions were incubated at 37°C during the assay. Enzyme activities
were determined spectrophotometrically at 340 nm in a Schimadzu UV-2401 PC
(Kyoto, Japan). The absorbance blank was a sample that contained water in place
of the cell-free cell extract.
Gluconokinase assay: Ten µl of cell- free extract were added to the
following final concentrations 50mM HEPES, pH 7.6, 10mM MgCl2, 3mM ATP, 1
mM potassium gluconate, 0.2mM NADP and 1 U 6-phosphogluconic
dehydrogenase. The final reaction volume was 1.0 ml. The reaction was started
by the addition of the 10 µl cell-free extract. All reactions were incubated at 37°C during the assay. Enzyme activities were determined spectrophotometrically at
340 nm in a Schimadzu UV-2401 PC (Kyoto, Japan). The absorbance blank was a sample that contained water in place of the cell-free cell extract.
Activity of each enzyme was measured as nmoles/min/mg of protein.
Enzyme assays were performed in duplicate. All samples were compared by
49 ANOVA using Tukey’s post-hoc test at the 95% confidence level (SigmaStat
version 3.1, Systat Software, Inc., Richmond, CA).
Gluconate uptake assay: Wild-type L. reuteri 100-23 and mutant L. reuteri
100-23D were grown to mid-log phase in MRS with glucose (100 mM), gluconate
(100 mM) or glucose + gluconate (50 mM gluconate and 50 mM glucose) as the
sole carbon source(s). Cells were centrifuged and washed in 0.05 M potassium
phosphate buffer, pH 6.8 with 0.2 g/L MgSO4. Cells were resuspended in 400 µl of the same buffer. 5 µg/ml of chloramphenicol were added to the cell suspension, and cells were incubated at 37°C for 5 minutes. 100 µl of cell
suspension were removed for protein analysis and 250 µl was placed into a new microcentrifuge tube. Gluconate uptake was started by the addition of 75 µl of radioactive gluconate (6mCi/mmol; 4.33 mM gluconate). Samples were taken at
0, 2, 5, and 20 minutes. Reactions were kept at 37°C. At each time point, 75 µl of the mixture was removed and washed by filtration three times with 5 ml of 250 mM gluconate prepared in 0.1 M potassium phosphate buffer, pH 6.0. The filter was placed into scintillation vials containing 10 ml of Scintiverse scintillation fluid (Fisher Scientific, Pittsburgh, PA). Radioactivity was quantified using a liquid scintillation counter 24 hours later (Beckman Coulter, Fullerton, CA). All samples were performed in duplicate.
50 The 100 µl of cell suspension removed prior to the uptake assay above was centrifuged. The cells were resuspended in 200 µl sterile water. A small amount of 0.1 mm glass beads was added, and the cells were pulverized 6 times for 30 seconds each with incubation on ice in between each 30 second treatment.
Samples were then centrifuged and resuspended in 200 µl of sterile water. Next,
10 µl of the suspension was added to 200 µl of diluted Bio-Rad dye reagent and analyzed as described above.
Construction of insertional mutation in gluconate genes of L. reuteri
Attempts to create a gluconate-negative mutant by ultraviolet light were
successful. However, an UV-generated mutant is not specifically mutated within
one gene and may possess other mutations. Therefore, a specific insertional
mutant was created. The method of Russell and Klaenhammer (48) was utilized.
First, the “helper” plasmid pTRK669 (RepA+, Cmr) was transformed into L.
reuteri 100-23 and the stability of pTRK669 was tested at 37°C and 43°C in the
absence of antibiotic selection. Transformation was performed by the method of
Luchansky et al (35, 36) with some modifications as described below. Cells (200
ml) were grown in MRS broth until late log phase (A600 = 0.8-1.0), harvested and
washed twice with 50 ml 3.5× SMEB (0.952 M sucrose; 3.5 M NaCl, pH 7.2). Cells
were then suspended in 10 ml 3.5× SMEB and held on ice for 10 minutes. One µg
51 of plasmid DNA was mixed with 400 µl cell suspension in a cold, sterile
microcentrifuge tube. The mixture was transferred to a 0.2 cm electroporation
cuvette and held on ice for > 1 minute. Samples were then electroporated in a
Gene Pulser (Bio-Rad Laboratories, Hercules, CA) at 2.5 kV; 200Ω, and 25µF.
Electroporated cells were then immediately added to 10 ml pre-warmed MRS
broth and incubated for 3 hours, anaerobically at 37°C. After incubation, cells
were centrifuged and resuspended in 0.3 ml of MRS and plated onto MRS with chloramphenicol (7.5 µg/ml ) agar plates.
PCR products internal to the putative gntK and gntP genes were each
ligated into pORI28 (Emr). Briefly, the putative gluconate kinase and permease fragments were subjected to PCR reactions in order to contain 5’ and 3’ ends compatible to SphI and PstI digests. The new gntK and gntP PCR fragments were then ligated into SphI and PstI digested pORI28. These constructs were transformed into E. coli EC1000 (RepA+) (30) by electroporation (49) and plated
onto LB containing kanamycin (40 µg/ml) and erythromycin (10 µg/ml) agar
plates (Figure 3.3). Transformants were analyzed to verify that they contained
plasmids with insertion of the PCR products in pORI28 (QIAprep Spin Miniprep
Kit; Qiagen, Valencia, CA). The resultant pORI28:gntK and pORI28:gntP
constructs and pTRK669 were then transformed into L. reuteri 100-23 in a two-
step process. First, pTRK669 was transformed into L. reuteri 100-23 (Figure 3.4).
52 Next, pORI28:gntK or pORI28:gntP constructs were transformed into L. reuteri containing pTRK669 using the transformation method described above (Figure
3.5 ). The transformed culture was then transferred daily in MRS with erythromycin (4 µg/ml) for 20 days at 48°C, anaerobic, to allow for loss of the
“helper” plasmid, pTRK669, and for homologous recombination of the pORI28- derived plasmid with the chromosome at the gene of interest (Figure 3.6). To identify colonies that had lost pTRK669 and acquired a chromosomal insertion of the pORI28-derived plasmid, colonies were spotted on MRS + erythromycin (4
µg/ml) and MRS + chloramphenicol plates. Chloramphenicol-sensitive, erythromycin-resistant colonies were plated on MRS and MRS-gluconate to confirm that loss of gluconate utilization coincided with insertion of the pORI- derived plasmid into gntK or gntP. To verify that disruption of the gluconate gene(s) was the result of integration of pORI28 harboring gntK and gntP fragments, Southern hybridization using StuI- and HpaI-digested genomic DNA and pORI28 and the gntK and gntP fragments as probes was performed.
53 pORI28 ori+ + gntK or gntP PCR product
Em
pORI28 ori+ PCR product repA Em chromosome
E. coli EC1000
Figure 3.3: Plasmid pORI28 ligated with putative gntK and gntP fragments transformed in E. coli EC1000.
54 ori+ pTRK669 repA
Cmr
gntP and gntK chromosome copy 37°C
ori+ pTRK669 repA
Cmr
L. reuteri 100-23
Figure 3.4: Transformation of pTRK669 into L. reuteri 100-23.
55 pORI28 ori+ PCR product
Emr
ori+ pTRK669 repA
37°C Cmr
pORI28 ori+ PCR product
Emr L. reuteri 100-23
Figure 3.5: Transformation of pORI28:gntP construct into L. reuteri 100-23 containing pTRK669.
56 48°C
Integrated pORI28
L. reuteri 100-23
ori+ pTRK669 repA When shift to 48°C is lost Cmr
Figure 3.6: Loss of pTRK669 and homologous recombination of pORI28:gntP with chromosomal gene copy when cells are upshifted to 48°C.
Transcriptional analysis
RNA extraction
Lactobacillus reuteri 100-23, mutant 100-23D, L. acidophilus NCFM and human isolates L rhamnosus 998 1A, L. rhamnosus 587 4B, L. paracasei subsp.
57 paracasei 59 5A and L. fermentum 59 5B were grown to an A600 of 0.3-0.6 in
chemically defined medium (29) and/or MRS containing either 100 mM glucose,
100 mM gluconate or glucose + gluconate (50 mM gluconate and 50 mM glucose). Isolation of RNA was performed using TRIzol as described by
Dinsmore and Klaenhammer (9). To isolate total RNA, cells (20 ml) were centrifuged and resuspended in 1 ml of TRIzol (Invitrogen, Carlsbad, CA). The
cell suspension was added to a small amount of 0.1 mm glass beads (BioSpec
Products, INC, Bartlesville, OK) and pulverized in a Mini Bead Beater-8 (BioSpec
Products, INC, Bartlesville, OK) for a total of 4 minutes in 1 minute intervals
with incubation on ice in between intervals. The cell extract was centrifuged for
10 minutes. The supernatant was then removed to a new tube and allowed to incubate at room temperature for 5 minutes. Next, 0.2 ml of chloroform was added. The tube was shaken vigorously for 15 seconds and incubated for 3 minutes at room temperature. Samples were then centrifuged for 15 minutes.
Following centrifugation, the upper phase was transferred to a new tube.
Isopropanol (0.5 ml) was added to the tube, and the mixture was incubated for 10 minutes at room temperature. The samples were then centrifuged for 10 minutes.
The RNA pellet was then washed with 75% ethanol prepared in DEPC-treated
water and dried (9). After drying, the cell pellet was resuspended in 30 µl DEPC-
treated water. The RNA concentration was determined spectrophotometrically at
260 nm in a Schimadzu UV-2401 PC (Kyoto, Japan).
58 To prepare RNA for loading on the gel, 40 µg of RNA was precipitated with 3 M sodium acetate, pH 5.2, and 2.5 volumes of ethanol. The RNA pellet was allowed to dry and then was suspended in 4.5 µl sterile DEPC-treated water.
To the RNA, 2.0 µl of 5× formaldehyde-gel-running buffer (0.1 M MOPS, pH 7.0;
40 mM sodium acetate; 5 mM EDTA, pH 8.0), 3.5 µl formaldehyde, and 10 µl of formamide were added. The RNA sample was then incubated at 65°C for 15 minutes. After incubation, samples were chilled immediately and 0.5 µl ethidium bromide (1% solution; Fisher Scientific, Pittsburgh, PA) and 2 µl gel-loading buffer (1 mM EDTA, pH 8.0; 0.25% bromophenol blue; 0.25% xylene cyanol FF prepared in 50:50 glycerol: DEPC-treated water) were added to each sample. A
0.24-9.5 Kb RNA ladder was used to estimate the transcript(s) size (Invitrogen,
Carlsbad, CA)
Gel preparation and electrophoresis conditions
The gel tray and casting unit were treated with RNase ZAP (Ambion Inc.,
Austin, TX). A 1% agarose gel was prepared in DEPC-treated water (62.25 ml), melted in the microwave and cooled. After cooling, 20 ml of 5× formaldehyde- gel-running buffer and 17.75 ml 37% formaldehyde were added. The gel solution was mixed and poured into the gel tray. After solidification, the gel was placed into the electrophoresis unit and covered with 1× formaldehyde-gel-running
59 buffer. RNA samples were loaded and electrophoresed at 80 V. RNA was visualized using a Bio-Rad digital imaging system (Bio-Rad Laboratories,
Hercules, CA).
Northern hybridization
The RNA-containing formaldehyde gel was washed three times (30 minutes each) in DEPC-treated water and one time in 20× SSC (49). Transfer of
RNA from the gel to a nylon membrane was performed by standard capillary transfer (49). After transferring over night, the nylon membrane was allowed to dry and cross-linked (UV Stratalinker 1800; Stratagene, La Jolla, CA). The membrane was then hybridized with digoxigenin-labeled putative gntP PCR product at various temperatures (38, 42 and 50°C) overnight. After hybridization, the membrane was then washed and subjected to reagents in the
DIG-high prime DNA labeling and detection starter kit II (Roche Applied
Sciences, Indianapolis, IN). Briefly, the membrane was washed at room temperature with various combinations and times of 0.5-5× SSC 0.1% SDS washes. Membranes were then subjected to reagents in the DIG-high prime DNA labeling and detection starter kit II (Roche Applied Sciences, Indianapolis, IN).
The membrane was then exposed to Kodak BioMax MS (Kodak, Rochester, NY) film and developed.
60 3.4. RESULTS AND DISCUSSION
Screening of bacteria for gluconate utilization
Results, shown in Table 3.2, indicate that all strains of L. zeae and L. reuteri use gluconate, whereas L. acidophilus and L. gasseri do not. This concurs with previous reports that L. zeae, closely related to L. casei, and L. reuteri use gluconate (2, 33). Four of the eleven P. freundenreichii subsp. shermanii strains were able to grow in minimal medium with gluconate as the sole carbon source.
Two of the three Bifidobacterium species tested were able to utilize gluconate. In contrast to the present study, previous studies (2, 70) found that B. adolescentis could utilize gluconate in broth and that B. breve and B. infantis could not utilize gluconate. Asano et al (2) also found that one strain P. acnes was capable of fermenting gluconate in broth. Lactobacillus reuteri 100-23 was chosen for further analysis due to its ability to utilize gluconate and to colonize the mouse intestine
(62)
Cloning of putative gluconate gene fragments from L. reuteri 100-23
Gluconate metabolism genes have not been characterized in Lactobacillus species; therefore, gene sequences from other bacterial genera were used to design degenerate primers (Table 3.1). A 750 base pair (bp) and 500 bp fragment were amplified using degenerate primer sets 3 and 6 (Table 3.1) based on
61 Carbon Source Carbon Source Strain (source) Gluconate Glucose Strain Gluconate Glucose P. freundenreichii subsp. L. reuteri 100-23 √ √ shermanii P2D √ (mouse intestine)a (cheese starter culture) P. freundenreichii subsp. L. reuteri ATCC 23272 √ √ shermanii P4D √ (human intestine) (cheese starter culture) P. freundenreichii subsp. L. gasseri ADH √ shermanii P5D √ (human intestine) (cheese starter culture) P. freundenreichii subsp. L. gasseri ATCC 33323 √ shermanii P6D √ √ (human intestine) (cheese starter culture) L. acidophilus ATCC P. freundenreichii subsp. 700396 √ shermanii P8D √ √ (human intestine) (cheese starter culture) P. freundenreichii subsp. L. acidophilus BG2FO4 √ shermanii PS31 √ (human intestine) (cheese starter culture) P. freundenreichii subsp. L. zeae ATCC 393 √ √ shermanii PS-1-A √ √ (cheese) (cheese starter culture) B. adolescentis ATCC P. freundenreichii subsp. 15703 √ shermanii PS-1-F ( √ (human intestine)b cheese starter culture) P. freundenreichii subsp. B. breve ATCC 15698 √ √ shermanii PS-4-B √ (infant intestine) (cheese starter culture) B. infantis ATCC P. freundenreichii subsp. 25962 (infant √ shermanii PS-5-H √ intestine) (cheese starter culture) P. freundenreichii subsp. shermanii P1D P. jensenii ATCC 4867 √ √ (cheese starter (buttermilk) culture)c a Cell growth measured in 20 g/L glucose or gluconate CDM broth b Cell growth measured 5 g/L glucose or gluconate reinforced clostridia broth c Cell growth measured in 12 g/L glucose or gluconate MMC d No growth observed in glucose or gluconate MMC
Table 3.2: Growth of selected Lactobacillus, Bifidobacterium and Propionibacterium strains from human or food origins with gluconate or glucose as the sole carbon source
62
gluconate kinase and permease sequences, respectively, as described in the
Materials & Methods section. Comparison of the amino acid sequences encoded
by the fragments showed that the 500 bp fragment had 47% sequence identity to
the gluconate kinase from Staphylococcus epidermidis and the 750 bp fragment had
53% sequence identity to the gluconate permease from C. acetobutylicum (June 10,
2003).
Hybridization of the L. reuteri 100-23 total genomic DNA to each of the
two labeled PCR fragments showed a single band of ~3 kb for both probes
(Figure 3.7). These results suggest that the putative gluconate kinase and
permease genes may be on the same DNA fragment, possibly arranged in an
operon. In B. subtilis, gluconate genes are arranged in an operon (~5.1 kb in
length), but transcribed as a polycistronic message (15). In E. coli, a 3.98 kb-
BamHI DNA fragment was found to contain a gluconate repressor (gntR), a
gluconokinase (gntK) and a permease (gntU) genes (66).
Colony hybridization
No positive clones were identified with colony hybridization experiments;
therefore, the complete gene sequence(s) involved in gluconate metabolism in L.
reuteri 100-23 could not be elucidated.
63
A B 1 2 3 4 1 2 3 4
Figure 3.7: Southern hybridization analysis of L. reuteri 100-23 using the putative gntK (A) or gntP (B) PCR product as the probe. Lane 1: putative gntK fragment (500 bp), Lane 2: putative gntP fragment (750 bp), Lane 3: blank, Lane 4: L. reuteri 100-23 genome digested with EcoRI.
Insertional mutagenesis of a putative gntP gene
An insertional mutagenesis strategy was employed to create site-directed
mutations in the putative gluconate metabolism genes of L. reuteri 100-23. As
Russell and Klaenhammer (48) found for L. acidophilus, the results indicate that
pTRK669, the helper RepA+ plasmid, is stable at 37°C but not at 43°C in L. reuteri, though more generations were required to cure the plasmid from L. reuteri 100-
23. A greater than seven-log reduction in chloramphenicol-resistant cells was
64 seen after daily culture transfer to fresh medium at 43°C for 20 days. No reduction was observed at 37°C.
Attempts to integrate the pORI28:putative gntK fragment plasmid into the
L. reuteri 100-23 genome were unsuccessful. No chloramphenicol-sensitive, erythromycin-resistant cells were recovered. However, after transferring L. reuteri 100-23 + pTRK669 culture transformed with pORI28:putative gntP fragment, for 20 days, chloramphenicol-sensitive, erythromycin-resistant colonies were recovered. Some of these colonies were also gluconate negative
To confirm that the gntP gene was disrupted, Southern analysis was performed on the putative insertional mutant genomic DNA cut with HpaI, which cuts the pORI28:putative gntP plasmid once. The putative gntP gene fragment or pORI28 were used as probes. Results shown in Figure 3.8 indicated that the gntP putative gene was disrupted by the recombination of pORI28:putative gntP fragment into the L. reuteri 100-23 genome in all three selected mutants. One mutant (D) was selected for further characterization and named L. reuteri 100-23D. To our knowledge, this is the first report of using insertional mutagenesis in L. reuteri.
Enzymatic activities associated with gluconate metabolism
To characterize which gluconate metabolism function(s) were defective in L. reuteri 100-23D, enzyme assays were performed on the wild-type and L. reuteri
65 100-23D (Tables 3.3 and 3.4). Gluconokinase activity was significantly higher in
cells grown in CDM with gluconate than MRS with gluconate grown cells.
Samples grown in glucose had virtually no gluconokinase activity in MRS or
CDM media. Gluconokinase activity in 100-23D was significantly lower than the wild-type when grown in MRS or CDM with equimolar glucose and gluconate.
When L. reuteri 100-23 and 100-23D were grown in MRS, there was no significant
difference in 6-phosphogluconate dehydrogenase activity. However, when cells
were grown in CDM, 6-phosphogluconate dehydrogenase activity was
significantly higher in the mutant, 100-23D, than the wild-type when grown in
equimolar glucose and gluconate. The low activity of 6-phosphogluconic acid
dehydratase suggests that L. reuteri 100-23 lacks this enzyme (data not shown).
Further studies are needed to verify this because the dehydratase enzyme is
unstable (69). Intracellular radioactive gluconate was significantly different between glucose-, gluconate- and glucose + gluconate-grown cells. These results
indicate that the mutant, 100-23D, has significantly reduced uptake and
gluconokinase activities. These results indicate that gluconate enzyme activities
are induced in the presence of gluconate and that gluconate genes in L. reuteri
100-23 are potentially arranged in an operon.
66
Strain Base Sugar Gluconokinase* 6phosphogluconate medium (µmol/min/mg of Dehydrogenase* protein) (µmol/min/mg of protein) L. reuteri 100-23 MRS glucose 0.0615f 0.131f L. reuteri 100-23 MRS gluconate 2.54b 0.149e,f L. reuteri 100-23 MRS glucose + 0.296d 0.142e,f gluconate L. reuteri 100-23D MRS glucose + 0.143e,f 0.185d,e gluconate L. reuteri 100-23 CDM glucose 0.149e,f 0.254c L. reuteri 100-23 CDM gluconate 3.73a 0.414a L. reuteri 100-23 CDM glucose + 0.548c 0.145e,f gluconate L. reuteri 100-23D CDM glucose + 0.182e 0.228c,d gluconate E. coli K12 CDM gluconate 0.040f 0.349b *Average values with the same letter within the same column are not statistically different (P>0.05), n=2.
Table 3.3: Enzyme activities of L. reuteri 100-23 and 100-23D
Growth medium Gluconate uptake rate (nmoles/min/mg protein) L. reuteri 100-23 MRS gluconate. 1.34a L. reuteri 100-23 MRS glucose + gluconate 0.232b L. reuteri 100-23D MRS glucose + gluconate 0.00805c *Average values with the same letter within the same column are not statistically different (P>0.05), n=2.
Table 3.4: Gluconate uptake rates for L. reuteri 100-23 and 100-23D
67 Transcriptional analysis
Transcription of the putative gluconate permease gene that was
interrupted by site-directed insertional mutagenesis was evaluated under
various growth conditions in L. reuteri 100-23 and 100-23D along with several
Lactobacillus strains isolated from human feces that are capable of utilizing
gluconate (see Chapter 2). Total RNA from the strains grown with different
sugars was hybridized to the 750 bp putative permease (gntP) PCR fragment.
Lactobacillus reuteri 100-23 and 100-23D had a single band which hybridized to
the gntP probe (Figure 3.9). This band was present when cultures were grown in
the presence of glucose, gluconate and glucose + gluconate mixture in MRS
broth. This indicates that the putative gntP gene in L. reuteri 100-23 is not
inducible at the transcriptional level. In contrast, enzyme activities results indicated that gluconate genes in L. reuteri 100-23 are inducible. Further studies
to elucidate the exact mechanism of gluconate metabolism in L. reuteri 100-23
need to be undertaken.
The five human Lactobacillus isolates screened did not possess RNA
hybridizing to the probe. Explanations for this include: (1) these strains may not
possess the gntP gene and uptake gluconate in another fashion, (2) the gntP gene
in these strains differs enough in nucleotide sequence from that of L. reuteri 100-
23 that the probe would not hybridize, or (3) the gntP gene was not expressed
under these conditions in these strains. These results were confirmed by
68 Southern analysis with the putative gntP product. Again, none of the five human isolates possessed any regions of similarity to the gntP gene product.
A B 1 2 3 4 5 6 7 8 1 2 3 4 5 6 7
Figure 3.8: StuI digests of L. reuteri 100-23 mutants A, B and D probed with putative gntP fragment (A) and pORI28 (B) as the probe. Lane 1: 100-23A StuI digested chromosomal DNA, Lane 2: 100-23B StuI digested chromosomal DNA, Lane 3: 100-23D StuI digested chromosomal DNA, Lane 4: L. reuteri 100-23 StuI digested chromosomal DNA, Lane 5: L. reuteri 100-23 with pTRK669 and pORI28:putative gntP StuI digested chromosomal DNA, Lane 6, Blank, Lane 7: pORI28, Lane 8: pORI28:putative gntP
69
A B 1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8 9
Figure 3.9: Northern analysis of L. reuteri 100-23 wild-type and 100-23D; RNA gel (A) probed with putative gntP (B) Lane 1: 100-23D grown in MRS glucose, Lane 2: 100-23D grown in glucose + gluconate MRS, Lane 3: blank, Lane 4: L. reuteri 100- 23 grown in glucose MRS, Lane 5: blank, Lane 6: L. reuteri 100-23 grown in gluconate MRS, Lane 7: blank, Lane 8: RNA ladder, Lane 9: putative gntP fragment.
Conclusion
Several lactobacilli, bifidobacteria and propionibacteria were screened for
gluconate fermentation. Of the bacteria examined, two bifidobacteria, three
lactobacilli and four propionibacteria were able to ferment gluconate. Of the
bacteria screened, Lactobacillus reuteri 100-23 was selected for further
characterization. Putative gntK and gntP gene fragments from L. reuteri 100-23
were found by PCR using degenerate primers. A gluconate-negative L. reuteri
100-23 was prepared via homologous recombination. To our knowledge, this is
70 the first report of insertional mutagenesis via homologous recombination in L. reuteri. The mutant, named L. reuteri 100-23D and the wild-type were characterized for gluconokinase, 6-phosphogluconate dehydrogenase and uptake enzyme activities. Gluconokinase and uptake activities were significantly higher in the wild-type than 100-23D. There was no significant difference in 6- phosphogluconate dehydrogenase between wild-type and 100-23D.
Transcriptional analysis indicated that gntP gene transcription is not inducible, but constitutively transcribed in the presence of glucose or gluconate.
Transcriptional analysis also revealed a single transcript. Overall, genetic characterization of gluconate metabolism genes in L. reuteri 100-23 was begun, but still needs further characterization.
71
APPENDIX A
EVALUATION OF A SWISS CHEESE MODEL SYSTEM
72 A.1. Abstract Aseptic Swiss cheese slurries were made with five different culture combinations from four different culture manufacturers. Slurries were compared to commercial cheeses prepared with the same culture combinations. Cheese and slurry time points were analyzed over time for pH, streptococci, lactobacilli and propionibacteria cell numbers, free amino acids and acetic, lactic, propionic and succinic acids. Slurries were sampled over a period of 48 hours which was to correspond to the warm room ripening period of commercial cheese samples.
The pH of the slurries started at 6.0 and dropped quickly to as low as 4.6, whereas the cheese pH started at 5.3 and increased slightly to 5.7. This drastic difference in pH could account for bacterial cell count differences between slurry and cheese, most notably the lack of Propionibacterium cell growth in the slurry.
All of the free amino acid measurements at all slurry time points were statistically different (P<0.05) from cheese warm room measurements. Most of the slurry organic acids (acetic, lactic, propionic and succinic) were statistically different from the cheese samples. Although the present model is not a good representation of Swiss cheese parameters, it is the first time an aseptic model has been utilized in Swiss cheese modeling.
A.2. Introduction
Swiss cheese is manufactured with using streptococci, lactobacilli and propionibacteria species. All three bacteria interact and play a vital role in Swiss
73 cheese flavor and body characteristics. In the U.S., Swiss cheese is typically ripened for 60-90 days. In Europe, the ripening period is much longer. This time frame makes it difficult to evaluate the success or failure of new culture combinations and/or manufacturing changes. Thus, development of a Swiss cheese model system that would allow rapid evaluation of new culture combinations or manufacturing parameters would be beneficial. Previously
Cheddar and Swiss slurry models (18, 27, 47, 53, 54) have been successful in reducing ripening time of cheeses. However, these systems were not prepared aseptically, lending themselves to growth of indigenous milk and environmental microflora. Therefore, it was the purpose of this paper to develop and evaluate an aseptic Swiss cheese slurry system to assess culture combinations and compare bacterial growth and flavor development of the slurry system to commercial Swiss cheeses.
A.3. Materials and Methods
Cultures
All Swiss cheese cultures were obtained from commercial culture companies and prepared per company’s recommendations. Five different culture combinations recommendations from four different manufacturers were used
(Table A.1).
74
Combination Company Streptococcus Lactobacillus Propionibacterium 2 A S787 L701 P745 5 B S884 L879 P835 8 C S392 L386 and L333 P318 9 C S392 L367 and L350 P318 10 D S141 L168 P183
Table A.1: Culture combination recommendations from four different culture suppliers used in the manufacture of Swiss cheese and Swiss slurries
Slurry preparation
Swiss cheese slurries were prepared aseptically in sterile miniature cheese
vats designed by Roberts et al. (47). One liter of fat-free UHT milk (Parmalat,
Grand Rapids, MI) was aseptically added to each sterile miniature vat. The milk
pH was adjusted to 6.0-6.1 using L-lactic acid (Sigma, St. Louis, MO). Sterile 10%
CaCl2 was then added to a final concentration of 0.02%. The milk was warmed to
33.3°C by placing the vat in a circulating water bath. After warming, 5.5 ml of a
2.5% chymosin solution prepared in sterile water (Chy-max extra, Chr. Hansen
Inc., Milwaukee, WI) was added to each vat. After the addition of chymosin, the starter cultures were added as per culture suppliers’ recommendations. Five
75 different cultures combinations were tested (Table A.1). Each combination was
used to prepare 2-4 replicate slurries. After culture addition and stirring, the vats
were placed back into the 33.3°C water bath. After 40-45 minutes, the curd was cut (1/2”) aseptically with a sterile wire cutter. The curd was then cooked by increasing the water bath temperature with intermittent stirring for 35-45
minutes until the curd and whey mixture reached 50°C. After cooking, the curd
was held at 50°C until the pH reached 5.8 at which time the whey was drained
through sterile cheese cloth. The curd was kept in the vat and placed back into
the water bath until the curd pH was 5.5-5.6. At this time, a sample was taken for
moisture analysis. Moisture was analyzed in the Microwave Moisture Solid
Analyzer: LabWave 9000 (CEM corporation, Matthews, NC) using the following
program: 60% power; 1.8-2.2 grams of slurry; 7 minute dry time. Sterile 22%
NaCl and water were added make the final slurry have 60% moisture and 1%
NaCl in the moisture phase. The curd was placed into a sterile Whirl-Pak® bag
(Nasco, Ft. Atkinson, WI) along with the water and salt and stomached for 5
minutes in a Seward Stomacher (Biomaster 80, Seward Co., Norfolk, UK). A
sterile pastry tip was placed in the bag and a corner was cut off of the bag to
allow the pastry tip to protrude. The cheese slurry was extruded through the tip
and aliquoted into sterile 15 ml Falcon tubes. Approximately 5 grams was
extruded into each tube. Tubes were then placed in an anaerobic jar with a gas-
76 pak (Becton, Dickinson and Company, Franklin Lakes, NJ) with the caps remaining loose. The anaerobic jar was then placed into a Forma Scientific anaerobe system (model 1025; 85% nitrogen, 10% hydrogen, 5% carbon dioxide), where the anaerobic jar was sealed. The anaerobic jar was then removed and incubated at 24°C. At 0, 8, 20, 32, and 48 hours, samples were removed for analysis. At each time point, anaerobic jars were removed from the incubator and placed in an anaerobic chamber, where the samples were removed and the jar re- sealed.
Cheese manufacture
Cheeses were manufactured in triplicate (except cheese 9 which was in duplicate) at Ragersville Cheese Company in Ragersville, Ohio, USA. The factory’s standard commercial cheese making procedure was used. This procedure includes a low cooking temperature (118-119°F) to allow for kosher- certified whey. Samples were taken before brining and at 30 (just out of the warm room), 60, 90 and 120 days of ripening. Cheese 10 analyses were only carried out through day 60.
Analysis
The pH of slurries and cheeses was measured at each time point with a spear-tip electrode and a Corning pH meter (Corning Incorporated Life Sciences,
77 Acton, MA) For bacterial analysis, cheese or slurry was serially diluted tenfold in
2% trisodium citrate, homogenized for 3 minutes and plated onto Rogosa SL
(lactobacilli) (Difco, Becton, Dickinson and Company, Franklin Lakes, NJ);
lithium glycerol agar (propionibacteria) (37) and M17 with 0.5% lactose and
0.15% lithium chloride (streptococci) (Difco, Becton, Dickinson and Company,
Franklin Lakes, NJ). Plates were incubated anaerobically (85% nitrogen, 10%
hydrogen, 5% carbon dioxide) at 30°C for 5-7 days (propionibacteria); 37°C anaerobically for 2 days (lactobacilli) and 42°C aerobically for 1 day
(streptococci).
Free amino acid concentration was determined by the cadmium ninhydrin microtiter plate assay (3) using L-leucine as the standard. Briefly, 2.5 grams of slurry or 5.0 grams of cheese were added to 25 or 50 ml of 50°C 125 mM trisodium citrate, respectively, and were shaken for 1 hour at 225 rpm in a benchtop shaker (New Brunswick Scientific, Edison, NJ) at 50°C. After shaking,
25 or 50 ml sterile water was added to slurry or cheese samples, respectively.
Samples were then serially two-fold diluted in 62.5 mM trisodium citrate and 100
µl of each dilution were added to 200 µl of water and 600 µl cadmium ninhydrin reagent (3). Samples were incubated at 84°C for 5 minutes. Samples were then
cooled to room temperature and centrifuged at 13,000 rpm for 5 minutes. 200 µl
of the supernatant were removed from each tube and placed into a microtiter
78 plate. Absorbance was read at 490 nm in a microtiter plate reader (Molecular
Devices, Sunnyvale, CA). A standard curve of known L-leucine concentrations
was used to calculate the free amino acid concentration of each sample. All
reactions were preformed in duplicate
Organic acids were detected using an Agilent 1100 HPLC equipped with a
multi-wavelength detector. Organic acids were analyzed using a Bio-Rad HPX-
87H column (Bio-Rad Laboratories, LaJolla, CA) with a 10 mN H2SO4 flow rate of
0.6 ml/min and a column temperature of 65°C. Organic acids were detected at
210 nm. Acetoin and propionic acids co-eluted in the same peak; however,
propionic acid does not absorb at 290 nm. Therefore, the propionic acid peak
area was calculated by subtracting the peak area at 290 nm from the peak area at
210 nm. Concentrations of organic acids were calculated using peak areas of
standard curves. Standard compounds were purchased from Fisher Scientific
(Pittsburgh, PA) and Sigma-Aldrich Chemical Company (St. Louis, MO).
Samples were prepared by the method of Judith Narvhus (personal
communication). Briefly, 1 gram of sample was weighed and extracted by the
addition of 8 ml acetonitrile (Fisher, Pittsburgh, PA) and 0.2 ml 1 N H2SO4.
Samples were mixed on a rotary shaker for 20 minutes and then centrifuged at 8,
000 rpm for 15 minutes in a RC-5 Sorvall centrifuge (Thermo Electron
79 Corporation, Asheville, NC). Supernatants were then filtered through a MSF-13
(Advantec MFS, Inc, Pleasanton, CA) filter and 20 µl was used for analysis.
Statistical analysis
Results from each analysis were averaged and compared between cheese
and slurry using ANOVA with Tukey’s t-test. Values with P<0.05 were
considered statistically different.
A.4. Results
For the purpose of this study, the 48 hour incubation of the slurry at 24°C
was intended to correspond with the 3-4 week warm room ripening phase (21-25
°C) in Swiss cheese manufacturing. The actual duration of slurry incubation that
would correspond to the end of warm room ripening was to be determined in
the study. The pH of the slurries started at 6.0 and dropped quickly to as low as
4.6 whereas the cheese pH started around 5.3 and increased slightly to 5.7 (Figure
A.1). Attempts to buffer the slurry system to reduce this drastic pH decrease
were unsuccessful (Limpisathian, unpublished results (32). This drastic
difference in pH could account for bacterial cell count differences between slurry
and cheese. For example, in combination 2, the propionibacteria cell number in
the slurry remained constant at 105 cfu/g whereas in the cheese, the
propionibacteria increased one log after warm room ripening (Figure A.2).
80 Without the growth of propionibacteria, many of the compounds contributing to
cheese flavor cannot be developed. Again in combinations 5, 8, 9 and 10, the propionibacteria did not grow (Figures A.3-A.6). All of the free amino acid measurements at all slurry time points were statistically different (P<0.05) from cheese warm room measurements (Figure A.7). Most of the slurry organic acids
(acetic, lactic, propionic and succinic) were statistically different from the cheese samples. In slurries 5, 8, 9 and 10, lactic acid increased dramatically between 0 and 48 hours. In contrast, lactic acid in the cheese combinations remained constant or decreased slightly (Figures A.9-A.12).
A.5. Discussion
The present Swiss cheese model system is not indicative of real Swiss cheese. Swiss cheese flavor development is dependent on the interactions of
streptococci, lactobacilli and propionibacteria starter strains. Complex
interactions occur making each culture combination responsible for a unique
flavor profile. All culture combinations utilized in the present study were recommended by culture manufacturers and some are currently used in the
Swiss cheese industry. In the present model, the growth of propionibacteria was
inhibited, lactic acid increased instead of decreasing, the pH decreased
dramatically, and free amino acids were present in small quantities. Singh and
Kristofferson (53) were able to develop Cheddar and Swiss cheese flavors in a
81 slurry system within a few days. In contrast to the Singh and Kristoffersen
study, the pH of the slurries in this study decreased dramatically. Efforts at
controlling the pH were unsuccessful. Propionibacteria are important for flavor
and eye development in Swiss cheese. This organism is affected by several
factors such as pH, NaCl content of the cheese, ripening temperatures, and other
starters, streptococci and lactobacilli proteolytic activity (28). Thierry et al (65) investigated the effect of lactobacilli on propionibacteria growth in an Emmental
“juice” medium. This study confirmed that the growth of propionibacteria is significantly affected by the growth and proteolysis of lactobacilli. The low pH is not an ideal growth medium for propionibacteria (28). In order for the present model to be successful, the pH must be controlled
As noted previously by Roberts and Wijesundera (47), UHT treated milk used in this experiment required a longer coagulation time (~40 minutes) and lowering of the milk pH ~6.0 with L-lactic acid.
The present model is not an ideal model to monitor new culture combination interactions or manufacturing variables due to the decrease in pH.
Attempts to control the pH were unsuccessful. However, with modifications, this aseptic Swiss slurry model might be helpful in determining new Swiss cheese cultures or processing parameters.
82
Days after cheese manufacture
0 20 40 60 80 100 120 140 6.2
6.0 Slurry 2 5.8 Slurry 5 5.6 Slurry 8 Slurry 9 5.4 Slurry 10 5.2 Cheese 2 pH Cheese 5 5.0 Cheese 8 Cheese 9 4.8 Cheese 10 4.6
4.4
4.2 0 50 100 150 200 250
Hours after slurry manufacture
Figure A.1: pH monitored over time for slurry and cheese combinations
83
Days after cheese manufacture
0 20406080100120140 10
9
8
7 cfu/g 6 Slurry streptococci Slurry lactobacilli 5 Slurry propionibacteria Cheese streptococci 4 Cheese lactobacilli Cheese propionibacteria
3 0 50 100 150 200 250 Hours after slurry manufacture
Figure A.2: Combination 2 bacterial cell counts
84
Days after cheese manufacture
0 20406080100120140 10
9
8
7
6 log cfu/g 5 Slurry streptococci Slurry lactobacilli 4 Slurry propionibacteria Cheese streptococci 3 Cheese lactobacilli Cheese propionibacteria 2 0 50 100 150 200 250 Hours after slurry manufacture
Figure A.3: Combination 5 bacterial cell counts
85
Days after cheese manufacture
0 20406080100120140 10
9
8
7 log cfu/g
6 slurry streptococci slurry lactobacilli 5 slurry propionibacteria cheese streptococci cheese lactobacilli cheese propionibacteria 4 0 50 100 150 200 250 Hours after slurry manufacture
Figure A.4: Combination 8 bacterial cell counts
86
Days after cheese manufacture
0 20 40 60 80 100 120 140 10
9
8
7
6 log cfu/g 5 slurry streptococci slurry lactobacilli 4 slurry propionibacteria cheese streptococci 3 cheese lactobacilli cheese propionibacteria 2 0 50 100 150 200 250 Hours after slurry manufacture
Figure A.5: Combination 9 bacterial cell counts
87
Days after cheese manufacture
0 20 40 60 80 100 120 140 10
9
8
7
6
log cfu/g log slurry streptococci 5 slurry lactobacilli slurry propionibacteria 4 cheese streptococci cheese lactobacilli 3 cheese propionibacteria 2 0 50 100 150 200 250 Hours after slurry manufacture
Figure A.6: Combination 10 bacterial cell counts
88
Days after cheese make
0 20 40 60 80 100 120 140 350
300 Slurry 2 250 Slurry 5 Slurry 8 200 Slurry 9 Slurry 10 150 Cheese 2 Cheese 5 100 Cheese 8 Cheese 9 50 Cheese 10
mmol Leucine/ kg cheese or slurry 0
0 50 100 150 200 250
Hours after slurry make
Figure A.7: Free amino acids over time for slurry and cheese combinations
89
Days after cheese manufacture
0 20 40 60 80 100 120 140 250
200 Acetic acid slurry Lactic acid slurry Succinic acid slurry 150 Acetic acid cheese Lactic acid cheese 100 Succinic acid cheese
50 mg/g of cheese or slurry
0
0 50 100 150 200 250
Hours after slurry manufacture
Figure A.8: Organic acid analysis of slurry and cheese combination 2
90
Days after cheese manufacture
0 20 40 60 80 100 120 140 60
50 Acetic acid slurry Lactic acid slurry Propionic acid slurry 40 Succinic acid slurry Acetic acid cheese 30 Lactic acid cheese Propionic acid cheese 20 Succinic acid cheese
10 mg/g ofmg/g cheese or slurry
0
-10 0 50 100 150 200 250
Hours after slurry manufacture
Figure A.9: Organic acid analysis of slurry and cheese combination 5
91
Days after cheese manufacture
0 20406080100120140 35
30 Acetic acid slurry Lactic acid slurry 25 Propionic acid slurry Succinic acid slurry 20 Acetic acid cheese Lactic acid cheese 15 Propionic acid cheese Succinic acid cheese 10
mg/g of cheese or slurry 5
0
-5 0 50 100 150 200 250
Hours after slurry manufacture
Figure A.10: Organic acid analysis of slurry and cheese for combination 8
92
Days after cheese manufacture
0 20406080100120140 80
Acetic acid slurry 60 Lactic acid slurry Propionic acid slurry Succinic acid slurry 40 Acetic acid cheese Lactic acid cheese Propionic acid cheese Succinic acid cheese 20 mg/g of cheese or slurry mg/g 0
0 50 100 150 200 250
Hours after slurry manufacture
Figure A.11: Organic acid analysis of slurry and cheese for combination 9
93
Days after cheese manufacture
0 10203040506070 30
Acetic acid slurry 25 Lactic acid slurry Propionic acid slurry 20 Succinic acid slurry Acetic acid cheese 15 Lactic acid cheese Propionic acid cheese Succinic acid cheese 10
5 mg/g of cheese or slurry
0
-5 0 20406080100120140
Hours after slurry manufacture
Figure A.12: Organic acid analysis of slurry and cheese for combination 10
94 LIST OF REFERENCES
1. Altschil, S. F., W. Gish, W. Miller, E. W. Myers, and D. J. Lipman. 1990. Basic local alignment search tool. Journal of Molecular Biology 215:403- 410.
2. Asano, T., K. Yuasa, K. Kunugita, T. Teraji, and T. Mitsuoka. 1994. Effects of gluconic acid on human faecal bacteria. Microbial ecology in health and disease 7:247-256.
3. Baer, A., I. Ryba, J. Meyer, and U. Butikofer. 1996. Microplate assay of free amino acids in Swiss cheeses. Lebensm.-Wiss. u-Technol. 29:58-62.
4. Bausch, C., N. Peekhaus, C. Utz, T. Blais, E. Murray, T. Lowary, and T. Conway. 1998. Sequence analysis of the GntII (subsidiary) system for gluconate metabolism reveals a novel pathway for L-idonic acid catabolism in Escherichia coli. J Bacteriol 180:3704-10.
5. Bernsmann, P., C. A. Alpert, M. J. Deutscher, and W. Hengstenberg. 1982. The bacterial PEP-dependent phosphotransferase system mechanism of gluconate phosphorylation in Streptococcus faecalis. FEBS Letters 138:101-103 . 6. Bolotin, A., P. Wincker, S. Mauger, O. Jaillon, K. Malarme, J. Weissenbach, S. D. Ehrlich, and A. Sorokin. 2001. The complete genome sequence of the lactic acid bacterium Lactococcus lactis ssp. lactis IL1403. Cold Spring Harbor Laboratory Press Genome Research:1-23.
7. Bougle, D., N. Roland, F. Lebeurrier, and P. Arhan. 1999. Effect of Propionibacteria supplmentation on fecal bifidobacteria and segmental colonic transit time in healthy human subjects. Scandinavian Journal of Gastroenterology 2.
95
8. Bradford, M. M. 1976. A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochem. 72:248-254.
9. Dinsmore, P. K., and T. R. Klaenhammer. 1997. Molecular characterization of a genomic region in a Lactococcus bacteriophage that is involved in its sensitivity to the phage defense mechanism AbiA. Journal of Bacteriology 179.
10. Drinan, F. D., and T. M. Cogan. 1992. Detection of propionibacteria in cheese. Journal of Dairy Research 59:1-5.
11. Entner, N., and M. Doudoroff. 1952. Glucose and gluconic acid oxidation of Pseudomonas saccharophilia. Journal of Biological Chemistry 196:852-62.
12. Esienberg, R. C., and W. J. Dobrogosz. 1967. Gluconate metabolism in Escherichia coli. Journal of Bacteriology 93:941-49.
13. Farber, J. M., and E. S. Idziak. 1982. Detection of glucose oxidation products in chillded fresh beef undergoing spoilage. Applied and Environmental Microbiology 44:521-524.
14. Fujita, T., and Y. Miwa. 1989. Identification of an operator sequence for the Bacillus subtilis gnt operon. Journal of Biological Chemistry 264:4201-6.
15. Fujita, Y., T. Fujita, Y. Miwa, J. Nihashi, and Y. Aratani. 1986. Organization and transcription of the gluconate operon, gnt, of Bacillus subtilis. The Journal of Biological Chemistry 261:13744-13753.
16. Gibson, G. R., and M. B. Roberfroid. 1995. Dietary modulation of the Human Colonic Microbiota: Introducing the Concept of Prebiotics. Journal of Nutrition 125:1401-1412.
17. Glatz, B. A., and K. I. Anderson. 1988. Isolation and characterization of mutants of Propionibacterium strains. Journal of dairy Science 71:769-776.
18. Harper, W. J., and T. Kristoffersen. 1970. Biochemical aspects of flavour development in Cheddar cheese slurries. J Agric Food Chem 18:563-566.
96 19. Hill, M. J. 1981. Diet and the Human Intestinal Bacterial Flora. Cancer Research 41:3778-3780.
20. Holzapfel, W. H., P. Haberer, J. Snel, U. Schillinger, and H. J. Veld. 1998. Overview of gut flora and probiotics. International Journal of Food Microbiology 41:85-101.
21. Huang, Y., and M. C. Adams. 2003. An in vitro model for investigating intestinal adhesion of potential dairy propionibacteria probiotic strains using cell line C2BBe1. Lett Appl Microbiol 36:213-6.
22. Isturiz, T., E. Palmero, and J. Vitelli-Flores. 1986. Mutations affecting gluconate catabolism in Escherichia coli. Genetic mapping of the locus for the thermosensitive gluconokinase. Journal of General Microbiology 132:3209-3212.
23. Jenkins, J. K., W. J. Harper, and P. D. Courtney. 2002. Genetic diversity in Swiss cheese starter cultures assessed by pulsed field gel electrophoresis and arbitrarily-primed PCR. Letters in Applied Microbiology 35:423-427.
24. Kimura, K., A. L. McCartney, M. A. McConnell, and G. W. Tannock. 1997. Analysis of fecal populations of bifidobacteria and lactobacilli and investigation of the immunological responses of their human hosts to the predominant strains. Appl Environ Microbiol 63:3394-8.
25. Kleessen, B., B. Sykura, H. J. Zunft, and M. Blaut. 1997. Effects of Inulin and Lactose on Fecal Microflora, Microbial Activity and Bowel Habit in Elderly Constipated Persons. American Journal of Clinical Nutrition 65:1397-1402.
26. Klemm, P., S. Tong, H. Nielsen, and T. Conway. 1996. The gntP gene of Escherichia coli involved in gluconate uptake. J Bacteriol 178:61-7.
27. Kristoffersen, T., E. M. Mikolajaik, and I. A. Gould. 1967. Cheddar cheese flavour. IV. Directed and accelerated ripening process. Journal of dairy Science 50:292-297.
28. Langsrud, T., G. W. Reinbold, and 1973. 1973. Flavor development and microbiology of Swiss cheese. A review. III. Ripening and flavor production. Journal of Milk and Food Technology 36:593-609.
97 29. Lauret, R., F. Morel-Deville, F. Berthier, M. Champomier-Verges, P. Postma, S. D. Ehrlich, and M. Zagorec. 1996. Carbohydrate utilization in Lactobacillus sake. Appl Environ Microbiol 62:1922-1927.
30. Law, J., G. Buist, A. Haandrikman, J. Kok, G. Venema, and K. Leenhouts. 1995. A system to generate chromosomal mutations in Lactococcus lactis which allows fast analysis of targeted genes. Journal of Bacteriology 177:7011-7018.
31. Li, X. Z., K. Poole, and H. Nikaido. 2003. Contributions of MexAB-OprM and an EmrE homolog to intrinsic resistance of Pseudomonas aeruginosa to aminoglycosides and dyes. Antimicrob Agents Chemother 47:27-33.
32. Limpisathian, P. unpublished results.
33. London, J. 1990. Uncommon pathways of metabolism among lactic acid bacteria. FEMS Microbiology Letters 87:103-112.
34. London, J., and N. M. Chace. 1979. Pentitol metabolism in Lactobacillis casei. Journal of Bacteriology 140:949-954.
35. Luchansky, J. B., E. G. Kleeman, R. R. Raya, and T. R. Klaenhammer. 1989. Genetic transfer systems for delivery of plasmid deoxyribonucleic acid to Lactobacillus acidophilus ADH: conjugation, electroporation, and transduction. J Dairy Sci 72:1408-17. 36. Luchansky, J. B., P. M. Muriana, and T. R. Klaenhammer. 1988. Application of electroporation for transfer of plasmid DNA to Lactobacillus, Lactococcus, Leuconostoc, Listeria, Pediococcus, Bacillus, Staphylococcus, Enterococcus and Propionibacterium. Mol Microbiol 2:637-46.
37. Madec, M.-N., A. Rauault, J. L. Maubois, and A. Thierry. 1996. Selective medium containing lithium and a polyol or antibiotic for counting propionibacteria. Official Gazette of the United States Patent and Trademark Office Patents 1192:1226.
38. Mallet, A. K., I. R. Rowland, and A. Wise. 1984. Dietary fat and cecal microbial activity in the rat. Nutrition and Cancer 6:86-91.
39. McCartney, A. L. 2002. Application of molecular biological methods for studying probiotics and the gut flora. Br J Nutr 88 Suppl 1:S29-37.
98
40. McCartney, A. L., W. Wenzhi, and G. Tannock. 1996. Molecular Analysis of the Composition of the Bifidobacterial and lactobacillus Microflora of Humans. Applied and Environmental Microbiology 62:4608-4613.
41. Muyzer, G., and K. Smalla. 1998. Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Antonie Van Leeuwenhoek 73:127-41.
42. Myhal, M. L., D. C. Laux, and P. S. Cohen. 1982. Relative colonizing abilities of human fecal and K-12 strains of Escherichia coli in the large intestine of streptomycin-treated mice. European Journal of Clinical Microbiology 1:186-192.
43. Peekhaus, N., and T. Conway. 1998. What's for dinner? Entner-Doudoroff metabolism in Escherichia coli. J Bacteriol 180:3495-502.
44. Perez, A., M. E. Nader de Macias, and G. Oliver. 1995. Propionibacteria in the gut: Effect on some metabolic activities of the host. Lait 75:435-45.
45. Ringø, E., H. R. Bendiksen, S. J. Gausen, A. Sundsfjord, and R. E. Olsen. 1998. The effect of dietary fatty acids on lactic acid bacteria associated with the epithelial mucosa and from faecalia of Artic charr, Salvelinus alpinus (L.). Journal of Applied Microbiology 85:855-864.
46. Roberfroid, M. B., J. A. E. Van Loo, and G. R. Gibson. 1998. The Bifidogenic Nature of Chicory Inulin and its Hydrolysis Products. Journal of Nutrition 128:11-19. 47. Roberts, M., and C. Wijesundera. 1995. Development of an aseptic cheese curd slurry system for cheese ripening studies. Australian Journal dairy Technology 50:66-69.
48. Russell, W. M., and T. R. Klaenhammer. 2001. Efficient system for directed integration into the Lactobacillus acidophilus and Lactobacillus gasseri chromosomes via homologous recombination. Appl Environ Microbiol 67:4361-4364.
49. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: A laboratory manual. Cold Spring Harbor Laboratory Press 2nd ed.:Cold Spring Harbor, N.Y.
99 50. Satokari, R. M., E. E. Vaughan, A. D. Akkermans, M. Saarela, and W. M. de Vos. 2001. Bifidobacterial diversity in human feces detected by genus- specific PCR and denaturing gradient gel electrophoresis. Appl Environ Microbiol 67:504-13.
51. Sears, H. J., H. Jones, R. Saloum, I. Brownlee, and F. Lamareaux. 1956. Persistence of individual strains of Escherichia coli in man and dog under varying conditions. Journal of Bacteriology 71:370-372.
52. Simpson, P. J., C. Stanton, G. F. Fitzgerald, and R. P. Ross. 2003. Genomic diversity and relatedness of bifidobacteria isolated from a porcine cecum. Journal of Bacteriology 185:2571-2581.
53. Singh, V. K., and T. Kristoffersen. 1970. Accelerated ripening of Swiss cheese curd. Journal of Dairy Science 52:349-354.
54. Sutherland, B. J. 1975. Rapidly ripened cheese curd slurries in process cheese manufacture. Australian Journal dairy Technology 30:138-142 . 55. Sweeney, N. J., P. Klemm, B. A. McCormick, E. Moller-Nielson, M. Utley, M. A. Schembri, D. C. Laux, and P. S. Cohen. 1996. The Escherchia coli K-12 gntP gene allows E. coli F-18 to occupy a distinct nutritional niche in the streptomycin-treated mouse large intestine. Infection and Immunity 64:3497-3503.
56. Sweeney, N. J., D. C. Laux, and P. S. Cohen. 1996. Escherichia coli F-18 and E. coli K-12 eda mutants do not colonize the streptomycin-treated mouse large intestine. Infect Immun 64:3504-11.
57. Tannock, G. 1995. Microecology of the Gastrointestinal Tract in Relation to Lactic Acid Bacteria. International Dairy Journal 5:1059-1070. 58. Tannock, G. 1995. Role of Probiotics, p. 257-269, Human Colonic bacteria Role in Nutrition, Physiology and Pathology. CRC Press.
59. Tannock, G., K. Munro, H. J. M. Harmsen, G. W. Welling, J. Smart, and P. K. Gopal. 2000. Analysis of the fecal microflora of human subjects consuming a probiotic product containing Lactobacillus rhamnosus DR20. Applied and Environmental Microbiology 66:2578-2588.
60. Tannock, G. W. 2001. Molecular assessment of intestinal microflora. Am J Clin Nutr 73:410S-414S.
100
61. Tannock, G. W., K. Munro, R. Bibiloni, M. A. Simon, P. Hargreaves, P. Gopal, H. Harmsen, and G. Welling. 2004. Impact of consumption of oligosaccharide-containing biscuits on the fecal microbiota of humans. Appl Environ Microbiol 70:2129-36.
62. Tannock, G. W., and D. C. Savage. 1985. Detections of plasmids in gastrointestinal strains of intestinal lactobacilli. Proc. Univ. Otago Med. Sch. 63:29-30.
63. Tannock, G. W., A. Tilsala-Timisjarvi, S. Rodtong, J. Ng, K. Munro, and T. Alatossava. 1999. Identification of Lactobacillus isolates from the gastrointestinal tract, silage, and yoghurt by 16S-23S rRNA gene intergenic spacer region sequence comparisons. Appl Environ Microbiol 65:4264-7.
64. Tetaud, E., S. Hanau, J. M. Welss, W. F. Le Page, M. J. Adams, S. Arkison, and M. P. Barrett. 1999. 6-phosphogluconate dehydrogenase from Lactococcus lactisL a role for arginie resudues in binding substrate and coenzyme. Biochem. Journal 338:55-60.
65. Thierry, A., D. Salvat-Brunaud, and J. L. Maubois. 1999. Influence of thermophilic lactic acid bacteria strains on propionibacteria growth and lactate consumption in an Emmental juice-like medium. Journal of Dairy Research 66:105-113.
66. Tong, S., A. Porco, T. Isturiz, and T. Conway. 1996. Cloning and molecular genetic characterization of the Escherichia coli gntR, gntK, and gntU genes of GntI, the main system for gluconate metabolism. J Bacteriol 178:3260-9.
67. Tuohy, K. M., S. Kolida, A. M. Lustenberger, and G. R. Gibson. 2001. The prebiotic effects of biscuits containing partially hydrolysed guar gum and fructo-oligosaccharides--a human volunteer study. Br J Nutr 86:341-8.
68. Walter, J., C. Hertel, G. W. Tannock, C. M. Lis, K. Munro, and W. P. Hammes. 2001. Detection of Lactobacillus, Pediococcus, Leuconostoc, and Weissella species in human feces by using group-specific PCR primers and denaturing gradient gel electrophoresis. Appl Environ Microbiol 67:2578-85.
101 69. Wang, I. N., and D. E. Dykhuizen. 2001. Variation of enzyme activities at a branched pathway involved in the utilization of gluconate in Escherichia coli. Evolution Int J Org Evolution 55:897-908.
70. Yaeshima, T., T. Fujisawa, and T. Mitsuoka. 1992. Bifidobacterium species expressing phenotypical similarity to Bifidobacterium adolescentis isolated from the feces of human adults. Bifidobacteria and Microflora 11:25-32.
71. Zablotny, R., and D. G. Fraenkel. 1967. Glucose and gluconate metabolism in a mutant of Escherichia coli lacking gluconate-6-phosphate dehydrase. J Bacteriol 93:1579-81.
72. Ziemer, C. J., and G. R. Gibson. 1998. An Overview of Probiotics, Prebiotics and Synbiotics in the Functional Food Concept: Perspectives and Future Strategies. International Dairy Journal 8:473-479.
102