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1989

Comparison of Clinical Isolates of KlebsiellaPneumoniae to Klebsiella Sp. Isolated FromSummer Harvested Eastern Oysters, CrassostreaVirginica (Gmelin).Thomas Elbert GrahamLouisiana State University and Agricultural & Mechanical College

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Recommended CitationGraham, Thomas Elbert, "Comparison of Clinical Isolates of Klebsiella Pneumoniae to Klebsiella Sp. Isolated From SummerHarvested Eastern Oysters, Crassostrea Virginica (Gmelin)." (1989). LSU Historical Dissertations and Theses. 4845.https://digitalcommons.lsu.edu/gradschool_disstheses/4845

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Order Number 9025306

Com parison o f clinical isolates o f Klebsiella pneumoniae to Klebsiella sp . iso lated from sum m er harvested eastern oysters, Crassostrea virginica (G m elin)

Graham, Thomas Elbert, Ph.D.

The Louisiana State University and Agricultural and Mechanical Col., 1989

Copyright © 1990 by G raham , T hom as E lbert. A ll rights reserved.

300 N. Zeeb Rd.Ann Arbor, MI 48106

Comparison of Clinical Isolates of Kiebsiella pneumoniae to Klebsiella sp. Isolated from

Summer Harvested Eastern Oysters, Crassostrea virginica (Gmelin)

A Dissertation

Submitted to the Graduate Faculty of the Louisiana State University and

Agricultural and Mechanical College in Partial Fulfillment of the

Requirements for the Degree of Doctor of Philosophy

inThe Department of Food Science

ByThomas Elbert Graham

M.S. Louisiana State University, 1983 B.S. Tulane University, 1978

December 1989

ACKNOWLEDGMENTS

The author is grateful to Dr. Auttis M. Mullins for providing financial support and for serving as his interim major advisor. Appreciation is also felt for Dr. Cameron R. Hackney for bringing the author into the Department of Food Science and for acting as his initial major advisor; and for Dr. Robert M. Grodner, Dr. Joseph A. Liuzzo and Dr. Samuel P. Meyers for their support and for serving on his committee. Also, appreciation is felt for Dr. Dudley D. Culley, Jr. and Dr. John M. Larkin for serving as his minor advisors. The author recognizes Diane Paille, Kathy Pierson (Technical Support Representative, Gilford Systems, Ciba Corning Diagnostic Corp.) and Dr. Larry Reily for their technical support. He thanks Dr. John M. Larkin for the use of equipment in the Department of Microbiology and Dr. Marc A. Cohn for the use of equipment in the Department of Plant Pathology. The author is grateful for the support of his family and friends, especially his mother and father, during his tenure at Louisiana State University. He thanks Dr. Jeffrey Smith for his assistance in reviewing this manuscript. Finally, the Author wishes to acknowledge Gulf and South Atlantic Fisheries Development Foundation, Inc. for the grant that made this work possible.

TABLE OF CONTENTSPage

Acknowledgements - . . iiTable of C o n t e n t s .......................................iiiList of T a b l e s ........................................... ivList of F i g u r e s ............................................ vA b s t r a c t ..................................................viIntroduction ............................................ 1Literature Review ..................................... 5

Oyster Industry ................................. 5Nature of the Pollution Problem ................. 8Bacterial Indicators and Shellfish ............. 124-Methylumbel1ifery1-B-d-glucuronide (MUG) . . 21Klebsiella........................................... 24

Materials & Methods ................................. 29

O r g a n i s m s ........................................... 29Isolation and Testing of Isolates ............. 29Deoxyribonucleic Acid (DNA) Isolation . . . . 35Moles % Guanine Plus C y t o s i n e ................... 39DNA-DNA H o m ology.................................. . 4 3

Results & Discussion ................................. 48Conclusions & Recommendations ....................... 72R e f e r e n c e s .............................................. 75V i t a ..................................................... 87

iii

LIST OF TABLES

TABLE PAGE

1. Organisms Used in This Investigation . . . 302. DNA Solutions Used in This Study................ 363. API 20E Test Results of This Investigation . 494. MAR of Selected Organisms Used in This Study 515. Fecal Coliform and MUG Reactions from This

Study .........................556. Moles % Guanine Plus Cytosine of the Strains

T e s t e d ......................................... 607. The Mol% G+C of Clinical Isolates Tested . . 618. The Mol% G+C of Oyster Isolates Tested . . .619. Levels of Homology of Organisms Examined . . 6310. Level of Homology of Clinical Isolates Tested 6411. Level of Homology of Oyster Isolates Tested . 6412. Recommended Modified Fecal Coliform Test . . 74

LIST OF FIGURES

FIGURE PAGE

1. API 20E Test Strip Identification Codes . . 332. Mol% G+C Denaturation Plot from a Gilford

2400 Spectrophotometer................... . 4 13. Mol56 G+C Denaturation Plot from a Gilford

Response II Spectrophotometer ............. 424. First Derivative Plot from a Gilford

Response II Spectrophotometer ............. 425. DNA Homology Plot from a Gilford 2400

Spectrophotometer . . . . 446. dA/Minute Determinations by a Gilford

Response 11 Spe c t r o p h o t o m e t e r ................467. Flow Chart for DNA A n a l y s i s ....................478. Percent DNA Homology for Taxonomic Groupings 669. DNA Homology Data (Second R u n ) ................ 67

v

ABSTRACT

Suspected environmental fecal coliform positive Klebsiella sp. oyster isolates were analyzed and compared to human clinical isolates of Klebsiella pneumoniae. Oyster isolates were urease negative (at 24 hours) and were identified as K. pneumoniae using API 20E biochemical test strips. The ability to produce fecal coliform reactions was lost by 6596 of the oyster isolates after undergoing several passages on brain-heart-infusion agar at 20° to 25°C. Tests with 4-methylumbelliferyl-B-d-glucuronide (MUG) found both oyster and clinical isolates to be MUG negative. Moles % guanine plus cytosine (mol96 G+C) information indicated that oyster isolates had mol96 G+C values similar to K_. pneumon iae, and DNA-DNA homology information indicated conclusively that chromosomal DNA from oyster isolates was indistinguishable from that of K_. pneumoniae, having homologies of >_6396 with a mean of 8296.

Key words: oyster, Crassostrea virginica, fecal coliforms,Klebsiella pneumoniae, urease negative, DNA, moles 96 G+C, DNA-DNA homology, 4-methylumbel1iferyl-B-d-glucuronide (MUG)

INTRODUCTION

Oysters are a popular and economically important seafood that have been widely consumed for many years. Louisiana is presently the largest producer of the American or Eastern oyster, Crassostrea virginica, in the United States.

Oysters, being filter feeders and growing in waters close to heavily populated areas, are closely monitored for possible contamination by human sewage, especially since they are sometimes eaten raw, or with minimal heat processing, and have been responsible for documented outbreaks of human enteric disease in the past.

The method presently employed to determine the sanitary quality of shellfish is the fecal coliform most-probable- number (MPN) test, which determines the level of organisms by statistical inference. Waters from which oysters are harvested and the oysters themselves are tested by this method. If fecal coliform levels exceed established limits, then harvest areas can be "closed" or "restricted" and any harvested oysters destroyed or confiscated.

During the summer of 1982, Louisiana oysters shipped to North Carolina were embargoed due to excessive fecal coliforms. Investigations indicated that the oysters were harvested from "approved" waters. Bacterial evaluations elucidated a situation which had been earlier identified but

1

never actively pursued, namely, that fecal coliforms during the summer months along the Gulf Coast often arepredominantly non-E^ coli fecal coliforms- Studies undertaken at Louisiana State University, Baton Rouge, LA showed that the harvested oysters, when taken from dockside and tested for fecal coliforms, were within legal limits but exceeded those limits after arriving at the receiving states. En route testing documented rapid increases in fecal coliform counts that were attributed to temperature abuse conditions.

Sacks of oysters are commonly held on the decks of oyster boats in the sun and heat and then stacked in refrigerated trucks such oysters in the bottom center of thestacks often do not reach the optimum shipping or holdingtemperature before the oysters reach their destination,mainly Florida, North Carolina and Virginia. Under such conditions, Klebsiella spp. multiplied rapidly, causing the high fecal coliform counts responsible for the embargoing of the Louisiana oysters.

Seasonal studies documented the prevalence of E. coli as the principal fecal coliform when temperatures in oyster growing waters were <22°C, whereas Klebsiella spp. predominated when water temperatures were warmer. These Klebsiella spp. were shown to have characteristics associated with organisms which are normally found in the estuarine environment, including the ability to grow at refrigeration temperatures, poor correlation with sewage bacteria, seasonal

variation, and little multiple antibiotic resistance.Because of this, it was speculated that these organisms were possibly a species of Klebsiella whose normal habitat is the estuarine environment, or possibly a subspecies of pneumoniae which could be differentiated from clinicalisolates of K. pneumoniae.

Interest in identifying these organisms stems from the fact that 1C. pneumoniae is a fecal coliform by definition, routinely isolated from human feces and is an opportunisticpathogen. Being such, these Klebsiella spp. being isolatedfrom oysters could be considered legitimate targets as indicator organisms along with E_. col i. However, if the Klebsiella spp. being isolated normally inhabit the estuarine environment, or are long term survivors, it could be argued that these Klebsiella spp. do not necessarily indicate the presence of human fecal material (or at least not recent fecal contamination) and should probably not pose a human health hazard. In this situation, Klebsiella spp. should only be taken into account in determining the general bacterial quality of the oysters.

Based on the hypothesis that the Klebsiella spp. being isolated from Eastern oysters harvested from along the Gulf Coast during the warmer months could be differentiated from human isolates of K_. pneumoniae, this investigation was undertaken to compare these Klebsiella spp. to clinical isolates of K. pneumoniae, using thermal denaturation and

renaturation of chromosomal deoxyribonucleic acid. The information gathered will be used to recommend modifying the procedure presently used to test the sanitary quality of oysters and to demonstrate if the thermal procedures have any practical value in the screening process.

LITERATURE REVIEW

Oyster Industry

Four species of oysters are of importance to the seafood industry in the United States (Burrell 1985), including the American or Eastern oyster, Crassostrea virginica; the Pacific oyster, C. gigas; the European oyster, Ostrea edulis; and the Olympia oyster, O. lurida♦ Of these, C. virginica accounts for 85% of the United States landings (Moody 1981). It is probably also the most studied oyster species (Burrell 1985). Its range in the United States is from Maine down the East Coast, along the Gulf Coast to Texas (Burrell 1985; Churchill 1920).

Oyster production (all species) in the United States is now approximately 60 million pounds annually, down from 175 million at the turn of the century (Burrell 1985; Moody 1981). The reduction is attributed to sustained harvests and the loss of habitat over the years (Cake 1982; Korringa 1976). Recently the harvest has been relatively consistent from year to year, although there continues to be a slow decline (Cake 1982). Gulf Coast production, however, has remained relatively stable since the 1940*3 (Dugas 1982; Korringa 1976). Louisiana harvests approximately 10 million pounds of C. virginica annually (Coleman 1985; Moody 1981).

The reason for Louisiana’s success is attributed to the

5

extensive sub-tidal beds along its coast created by the action of the Mississippi, Atchafalaya, Pearl, and Sabine rivers (Coleman 1985; Korringa 1976). In these areas, the salinities are high enough to allow the oysters to grow but low enough to minimize their disease and predator problems (Burrell 1985; Coleman 1985; Dugas et al. 1982; Korringa 1976).

Methods for harvesting oysters are labor intensive and have remained virtually unchanged since the beginning of this century (Dugas et al. 1982; Churchill 1920). Two methods are used, tonging and dredging (Moody 1981; Churchill 1920). Along the Gulf Coast, dredging with motorized vessels is used almost exclusively (Cake 1982; Dugas et al. 1982; Korringa 1976). Once removed from the water, oysters are culled by hand to break up clumps of multiple oysters and to remove empty shells. Live oysters are generally piled onto the decks of the oyster boats, and time permitting, or at the end of the harvest period, one and a half bushels are placed into burlap sacks (Coleman 1985; Korringa 1976). The sacks (placed in stacks) remain on the deck until the boat returns to the dock for unloading at the end of the harvest period. By law, each sack must be tagged to indicate the individual who owns the lease where the oysters were taken and the location where the oysters were taken (NSSP 1988; Moody 1981). At the dock, if the oysters are scheduled to reach their destination within four hours, the sacks are placed on

open flat bed trucks for transportation. If the oysters are being shipped for longer periods, refrigerated (0° to 4°C) trucks are used (NSSP 1988). Regulations require thatvehicles be prechilled and that there be air channels or pallets under the sacks of oysters. The regulations further require that a temperature recording device be used. Temperatures must not remain above 10°C for over 12 consecutive hours.

8Nature of the Pollution Problem

With oysters being filter feeders, there is considerable concern for the sanitary condition of harvested oysters, as they are often eaten raw or with minimal cooking (Hunt et al. 1984; Wood 1976). Oysters from polluted waters have been implicated in past outbreaks of enteric disease (NSSP 1988; Dugas et al. 1982; Wood 1976).

Under optimal conditions an oyster can filter up to 100 gallons of water daily, and retain any ingested bacterial pollutants from the filtered water (NSSP 1988; Coleman 1985; Dugas et al. 1982; Perkins et al. 1980; Cabelli & Heffernan 1970). Therefore, monitoring of the growing waters and the oysters for possible presence of fecal contamination is required (NSSP 1988). The monitoring has been responsible for the closure of many oyster beds, thereby making the harvesting of oysters from those beds illegal. At any one time in Louisiana it is not uncommon for 25% to 30% of all beds to be closed to harvesting (Dugas et al. 1982). Therefore, creating a secondary problem of illegally harvested oysters (Cake 1982; Dugas et al. 1982).

Coastal erosion, due in part to the channeling of rivers and industrial endeavors (mainly the production of petroleum) has allowed the intrusion of salt water inland (Coleman 1985; Dugas et al. 1982; Korringa 1976). Also, as populations increase and wetlands are destroyed, the oyster beds are

brought into closer proximity to populated areas. Therefore, pollution problems are likely to increase. These problems are due, in part, to reliance on aging and over-burdened sewage treatment facilities that allow raw or partially treated sewage to be released into open waterways (Cake 1982; Dugas et al. 1982; Wood 1976). Also, there are a number of small communities with little or no sewage treatment. Industrial facilities (mainly petroleum related) and recreational fishing camps, without adequate sewage handling, further add to this problem.

Human fecal pollution has been estimated at 1.6x10’' bacteria per day per capita (NSSP 1988). The report goes on to state that it requires 8 million ft3 of unpolluted water to dilute the above dump load to an acceptable level (< 70 total coliforms per 100 ml). Compounding the problem is pasture runoff due to rain, which places warm blooded animal fecal material directly into the water with no treatment whatsoever (Coleman 1985; APHA 1984; Eyles & Davey 1984; Dugas et al. 1982; Hunt 1980; Wood 1976). All the above iscompounded along the Gulf Coast during the summer months by southerly winds which may reduce water exchange in estuaries, resulting in oyster growing waters becoming stagnant and eutrophic (Paille et al. 1987; Wood 1976).

Improper handling of harvested oysters further complicates the pollution problem (Wentz et al. 1983; Wood 1976). As mentioned above, once the oysters are removed from

the water, they are left on the decks of the oyster boats for considerable lengths of time. During the warmer months, withexposure to the sun and air temperatures at or above 90°F(32°C) for hours, there is a rapid proliferation of bacteria within these oysters (Paille et al. 1987; Reily et al. 1985;Sbaih et al. 1983). Oysters transported short distances tolocal processing plants often are not cooled sufficiently to allow the oysters to reach an optimum holding temperature (0° to 4°C) until within the plants. Studies have shown thatunder the above conditions, even when using proper handling procedures, bacterial levels can rise (Reily et al. 1985; Kelly 1961). Even if the oysters are shipped using refrigerated trucks, because of the way they are stacked in the trucks, some of the oysters may not reach the required shipping temperature (0° to 4°C) before the oysters reach their destination. Seven to eighteen hour transit times may not allow the sacks in the center of the stacks to reach the temperature at which the refrigeration unit is set. Temperature abuse in these cases is a reality and causes bacterial quality problems (Sbaih et al. 1983).

Concerns over bacterial contamination could be lessened by using purification methods such as depuration or relaying (Richards 1988; Cook & Ellender 1986; Eyles & Davey 1984; Rowse & Fleet 1984; Perkins et al. 1980; Hartland & Timoney 1979; Wood 1976). Using these methods, oysters from marginal or restricted beds are transported to and depurated, under

controlled conditions for 48 hours, in specially designed on­shore plants, or are transported (relayed) to "clean" waters, and allowed to purge for 14 days under ideal conditions, then harvested for human consumption (NSSP 1988).

12Bacterial Indicators and Shellfish

Because of the number of infectious bacteria and theiroften specialized growth requirements, examination of samplesfor specific disease organisms is not generally carried outunless one or more of these bacteria are suspected of beingpresent. It has become apparent over the years that certain disease organisms can, for the most part, be found in thepresence of other bacteria (Mehlman 1984). These bacteriaact as indicators for the possible presence of diseaseorganisms.

In order for organisms to be considered appropriate indicators, they must meet certain criteria (Sbaih et al. 1983; Banwart 1981):

1) They must be found in the presence of the pathogen(s ).

2) They must be associated with feces.3) They should not be present in the absence of feces.4) They must occur in greater numbers than the

pathogen(s).5) Their survival should reflect that of the

pathogen(s ).6) They should not reproduce in the estuarine

environment.7) They should be easy to grow and differentiate.

With regards to the sanitary quality of shellfish, the presence of indicators resulting from sewage contamination is the concern. The degree of sewage contamination in receiving waters depends on both physical and biological factors (Wood1976). The physical factors include the location of the

discharge point of the sewage, the quantity and concentration of the sewage effluent, the quality of the sewage effluent, and the dilution and dispersal characteristics of the receiving waters. The biological factors include thebacterial content of the sewage, the viability of the sewage microorganisms in the receiving water, and the behavior of the shellfish of concern.

Surveys of the history of bacterial sanitation quality indicate that the original indicator of sewage contamination was E_. col 1 (APHA 1984; Mehlman 1984; Banwart 1981). Since it can take up to 10 days to confirm the presence of this organism, using standard American Public Health Association (APHA) approved methods, other indicators were sought. A replacement for E. coli came with the institution of coliforms as the indicators of choice in 1914 (Mehlman 1984). Routine testing of shellfish was not started until 1925. This involved the use of the coliform test as the indicator of their sanitary condition (NSSP 1988).

Coliforms, by definition, are those bacteria that are gram negative, rod shaped, facultative anaerobes which are capable of fermenting lactose to produce acid and gas at 35°C in 48 hours (APHA 1984, Mehlman 1984, Banwart 1981). The coliform group includes coli, Erwinia sp., Enterobacteraerogenes, E_. cloacae, Citrobacter fruendii and Klebsiella spp. (APHA 1984). Coliforms were used to identify the possible presence of fecal material in shellfish and the

waters from which they are harvested. As time progressed it became apparent that there was a problem in using the so called "total" coliforms to indicate the presence of possible human sewage (Mehlman 1984; Blair & Michener 1961; Kelly 1961). It was found that total coliforms as a group include organisms which are naturally present in the estuarine environment and which belong to animals whose natural bacterial flora pose no direct health threat to humans. Also, it was found that certain non-coliforms recovered from the environment mimicked coliforms and interfered with the accuracy of the test (NSSP 1988; Sbaih et al. 1983; Austin et al. 1981; Hussong et al. 1980; Neilson 1978). After much debate, the indicator of choice for shellfish was changed in 1974 to fecal coliforms (NSSP 1988).

Fecal coliforms, by definition, are those bacteria that are gram negative, rod shaped, facultative anaerobes that can ferment lactose to produce acid and gas at 44.5° or 45.5°C in 24 hours (NSSP 1988; APHA 1984; FDA 1984; Mehlman 1984; Banwart 1981). The use of these organisms as the indicator of choice with regards to shellfish in the United States persists today. However, several European countries,including Belgium, Denmark, England, France, Italy, and The Netherlands use E. coli as their specific indicator (Dufour 1977; Wood 1976).

The validity of the fecal coliform test rests on the evidence that >90% of the fecal coliforms isolated by the

test are E. coli (Feachem et al. 1983; Hood et al. 1983a; Lavoie 1983; Banwart 1981; Dufour 1977; Tennant et al. 1961). This is because E. coli represents the greatest proportion of organisms excreted in the human feces (Lavoie 1983; Dufour1977). Also, it has been shown that a positive relationship exists between human sewage contamination in shellfishgrowing areas and human enteric disease outbreaks (NSSP 1988). It should be noted, however, that the terms "fecal coliform" and "coliform" have no taxonomic validity, since bacteria not normally associated with feces can fit eachdefinition (APHA 1984).

The fecal coliform test as it is now used, with regard to shellfish testing, is a 72 hour test (APHA 1984). Samples are prepared as per APHA approved methods, inoculated into lauryl tryptose (LST) broth using a five tube most-probable- number (MPN) method, and incubated for 24 hours at 35°C. At 24 hours, organisms from positive tubes (those tubes with growth and gas production) are serially transferred to tubes of EC medium. Negative LST tubes are reincubated as above for an additional 24 hours. EC tubes are incubated at 44.5°C in coliform incubator baths. After 24 hours, positive ECtubes (those with growth and gas) are recorded, with organisms from any additional positive LST tubes inoculated into EC medium, and incubated as above. After 72 hours, a final MPN reading is made, and a fecal coliform value determined using MPN charts (APHA 1984; FDA 1984). These

values are a statistical interpretation of the numbers of organisms it would take to make the test results conform to predetermined standards.

Confirming the presence of E_. col i, requires additionaltests (APHA 1984; FDA 1984). Levine eosine methylene blue(L-EMB) agar streak plates are used to make a presumptiveidentification of E_. col i. Growth from EC medium positivetubes are streaked to L-EMB for isolation and the plates incubated at 35° C. After 24 hours, presumptive E. col i isidentified by looking for small flat colonies with eitherdark centers or those which have a metallic green sheen.Colonies showing typical E_. col i morphologies are streakedonto nutrient agar (NA) slants for further testing.Confirmation of E_. col i requires an additional set ofbiochemical tests, IMViC for short, to be run. The IMViCreactions involve tests for indole production (24 hours), pHchange using methyl red (96 hours), Voges-Proskauer (48hours) reaction and citrate utilization (96 hours).Therefore, confirming E_. col i, requires 72 hours for thefecal coliform test, 24 hours for L-EMB, 24 hours on NA, and96 hours for the IMViC tests, for a total of 216 hours or 9days (FDA 1984). This is the reason, based on the fecalcoliform test identifying 95% E coli, that testing isstopped at 72 hours.

In order for growing waters to be approved, fecal coliform (FC) values of samples taken from those waters must

not exceed 14 FC/100 ml, with no more than 10% of the samples tested exceeding 43 FC/100 ml. In order for shellfish to be considered safe for human consumption values must not exceed 230 FC/100 gm, with no more than 10% of the samples tested exceeding 330 FC/100 gm (NSSP 1988; APHA 1984; Cockey 1983).Samples with FC values in excess of those mentioned above are considered to be out of compliance. Excess fecal coliform levels in shellfish growing waters will cause those waters to be "restricted" or "closed" to harvesting. Shellfish samples with excess fecal coliforms are considered adulterated and unfit for human consumption.

The use of fecal coliforms as the indicators of choice has been under debate for some time. Some people, however, feel that the fecal coliform test is suitable as presently used. Their reasons include that excess fecal coliform concentrations are due to improper handling (Wentz et al. 1983) ; that the test accurately reflects the level of E. coli, or human pollution (Cook & El lender 1986; Hood et al. 1983a; Tennant et al. 1961); or that the detection of non-E. coli fecal coliforms, such as Klebsiella and Enterobacter, by the test, still may indicate human pollution or potential problems, and do not invalidate the test (Woodward et al. 1979; Knittel et al. 1977; Knittel 1975). Both Klebsiellaand Enterobacter have been routinely isolated from human feces (Lavoie 1983, Dufour 1977).

Generally, the above authors contend that the test does

indicate true levels of fecal contamination. Furthermore, they contend that looking for E_. col i directly may sacrifice the test's reliability, even if some or many of the fecal coliforms being identified are not E. coli. Hunt (1980), however, while finding no constant level of pathogens associated with human sewage, feels that there is no better substitute for the fecal coliform test.

On the other hand, it has been known for some time thatfecal coliform testing does not always reflect accuratelevels of E_. col i or at least recent fecal contamination (Andrews et al. 1987; Stanetz et al. 1968). Along the Gulfcoast during the summer months, and in other areas where water temperatures are elevated, Klebsiella and Enterobacter can account for a large percentage of the fecal coliforms isolated by the test (Paille et al. 1987; Lopez-Torres et al. 1987; Joncas et al. 1985; Caplenas & Kanarek 1984; Hood etal. 1983b; Sbaih et al. 1983; Hussong et al. 1981; Casper1974; Adams 1972). Paille et al. (1987) indicated that fecalcoliforms isolated over the course of a year from oysters and their growing waters showed seasonal variations. Somesamples, taken during the summer months, had Klebsiella spp. as the only fecal coliforms recovered. Even though K_.pneumoniae is present in human feces, reports indicate that Klebsiella only makes up about 1% of the fecal bacteria and 12% of the fecal coliforms (Lavoie 1983; Dufour 1977).

Poor recovery of E_. col i, compared with other fecal

coliforms or interfering organisms, may be due to adverse conditions in the estuarine environments. High temperatures have been shown to reduce the survival of E. coli (Rhodes & Kator 1988; Flint 1987; Paille et al. 1987; Caplenas & Kanarek 1984; Hiraishi et al. 1984; Anderson et al. 1983;Sbaih et al. 1983; McCambridge 8k McMeekin 1981; Vasconcellos& Swartz 1976; Tennant et al. 1961). Problems due to high temperatures, with regards to the fecal coliform test, have been shown iji vitro. Samples grown at 44.5° and 45.5°C exhibited different relative growth rates of different coliforms (Caplenas & Kanarek 1984; Dutka 1979; Fishbein 8k Surkiewicz 1964; Tennant 8k Reid 1961). The 44.5°C tests resulted in up to 40% false positives, while the 45.5°C testsshowed only 10% false positives.

Solar radiation and bacterial predators have also been implicated in the reduced survivability of E_. coli (NSSP 1988; Rhodes 8k Kator 1988; Flint 1987; McCambridge 8k McMeekin 1981; McCambridge & McMeekin 1980). Yet another possible reason for reduced survivability and recovery of E_. col i may be due to sensitivity to salt (Roth et al. 1988).

The presence of fecal coliforms in unpolluted or pristine areas has also led to suspicion of the fecal coliform test as an indicator of human fecal pollution (Riviera et al. 1988; Adams 1972; Duncan 8k Razzel 1972; Stanetz et al. 1968).

Finally, there are those who doubt that the presence of

20fecal coliforms represents severe health threats to humans (Krumperman 1983; Brown & Seidler 1973).

4-Methylumbelliferyl-B-d-glucuronide (MUG)21

The MUG reagent has been known for some time (Mead et al. 1955). This reagent’s usefulness is based upon the ability of the enzyme B-glucuronidase to split MUG to produce the fluorescent compound (4-methylumbelliferyl), which can be detected using long wave ultraviolet light (Stratman 1988). It has been documented that >90% of E_. col i produce this enzyme while most of the other fecal coliforms do not, or do so at a much lower frequency (Iritani & Inzana 1988; Andrews et al. 1987; Rippey et al. 1987; Perez et al. 1986; Robinson1984; Feng & Hartman 1982; Kilian & Bulow 1976). Investigations incorporating MUG into the fecal coliform test have shown that it is effective in detecting E_. col i. One study indicated that it was 100% effective in detecting entcropathogenic E_. coll (Feng & Hartman 1982). In general, however, MUG is credited with a >95% efficiency rate in detecting E_. col i. Some studies even indicate that tests that incorporate the MUG reagent are actually more sensitive in detecting E. col i than those without (Rippey et al. 1987; Moberg 1985). It has been suggested that MUG can detect low concentrations of E_. col i , even when its presence is masked by the growth of other organisms. In addition, E_. coli may be detected when present in concentrations below the detection threshold of the standard reagent (Rippey et al. 1987; Feng & Hartman 1982).

False positives occur with some of the other enterobacteriaceae which can also produce B-glucuronidase. These include Salmonella, Shigella and Citrobacter (Perez et al. 1986). The ability to produce the enzyme, however, is not ubiquitous in these populations. In general, it has been reported that only about 20% to 30% of Salmonella and Shigella are capable of producing the enzyme (Perez et al. 1986; Feng & Hartman 1982; Le Minor 1979).

The usefulness of MUG is that it can detect E. coli while excluding other fecal coliforms, especially Klebsiella and Enterobacter. With regards to both environmental and clinical isolates of Klebsiella, reports indicate negative results with MUG reagent, i.e., no fluorescence is produced (Perez et al. 1986; Feng & Hartman 1982; Kilian & Bulow 1976). The reason .is that Klebsiella does not produce B- glucuronidase. In one study, however, one tube whichproduced a fluorescent reaction yielded no E. coli but did showed Klebsiella (Koburger & Miller 1985). The authors speculated that a synergistic interaction may have occurred to produce the positive results, since a retest with the recovered Klebsiella did not produce a fluorescent reaction.

With regard to shellfish, there is a problem with MUG. B-glucuronidase produced naturally in shellfish tissues causes false positive reactions in fecal coliform-MUG tests (Watkins et al. 1988; Poelma et al. 1987; Rippey et al. 1987; Koburger & Miller 1985). In these situations MUG was

incorporated into the LST broth, where high levels of tissues were added when inoculating the media. This situation was solved by incorporating MUG into the EC medium. When this was done, false positives due to the shellfish enzyme were not detected (Rippey et al. 1987; Koburger & Miller 1985).

24Klebsiella

Information on Klebsiella abounds in the literature. Paille et al. (1987) tested oyster samples which yielded what appeared to be almost pure cultures of Klebsiella sp. They speculated that these Klebsiella sp. might be species whose normal habitat is the estuarine environment, since they possessed environmental characteristics. Thesecharacteristics included the ability to grow at refrigerator temperatures, seasonal variation, poor correlation to sewage, and little multiple antibiotic resistance (MAR). Strict environmental species of Klebsiella do exist and include K. terr igena (Izard et al . 1981) and K_. plant icola (Bagley etal. 1981). Also, K_. pneumoniae has routinely been recoveredfrom the environment (Richards 1988; Riviera et al. 1988; Lopez-Torres et al. 1987; Paille et al. 1987; Hiraishi et al.1984; Hood et al. 1983b; Lavoie 1983; Sbaih 1983; Evans etal. 1981; McCambridge & McMeekin 1981; Woodward et al. 1979;Bagley a Seidler 1977; Clark a Pagel 1977; Vlassoff 1977; Campbell et al. 1976; Knittel 1975; Casper 1974; Matsen et al. 1974; Duncan a Razzel 1972; Nunez a Colmer 1968).Klebsiella pneumoniae appears to have enhanced survivability, as compared to E_. col i, in the environment, even when from non-environmental sources, and can maintain its MAR for some time (Talbot et al. 1980; Knittel et al. 1977). There hasbeen some speculation that part of the abundance of

Klebsiella sp- is due to maintenance and possibly regrowth in the environment (Lopez-Torres et al. 1987; Hiraishi et al. 1984; Vasconcellos & Swartz 1976). Klebsiella is routinely isolated from plant material which may act as a reservoir for the organisms in the environment (King et al. 1988; Knittel et al. 1977; Seidler et al- 1977; Knittel 1975; Duncan ft Razzel 1972; Nunez ft Colmer 1968).

A possible reservoir for Klebsiella sp. in the estuarine environment may be the sediment (Paille et al. 1987). Klebsiella is found in higher concentrations in the sediment than in the overlying water. Klebsiella have the ability to fix nitrogen (Jones 1982; Knittel et al. 1977; Seidler et al. 1977; Brown & Seidler 1973; Nunez ft Colmer 1968). Studies have shown that estuarine sediments tend to exhibit a high degree of nitrogen fixation when temperatures are at or above 30° to 35°C (Jones 1982, Teal et al. 1979; Zuberer & Silver1978). Klebsiella have been isolated from these sediments.

There has been some disagreement as to how extensively Klebsiella is found in the environment. Some claim it is ubiquitous in nature (Paille et al. 1987; APHA 1984; Bagley ft Seidler 1977; Matsen et al. 1974). One investigator (Knittel 1975) disagrees and reports that in situations where it is found, E. coli is also isolated. Others have isolated Klebsiella in the absence of E_. coli (Paille et al. 1987; Sbaih et al. 1983; Seidler et al. 1977).

Klebsiella isolates from the environment have been

identified as being biochemically similar or identical to clinical isolates of K. pneumoniae (Paille et al. 1987; Caplenas ft Kanarek 1984; Bagley & Seidler 1977; Seidler et al. 1975; Brown 8k Seidler 1973). One difference which has been seen between the environmental and clinical strains is the inability or reduced ability of some environmental isolates to hydrolyze urea (Paille et al. 1987; Naemura et al. 1979). Paille et al. (1987) found that 69% of theirKlebsiella sp. isolates from oysters possessed this characteristic. Other investigators, however, have reported their environmental isolates as being able to hydrolyze urea (Seidler et al. 1975; Matsen et al. 1974). There was also an apparent reduced ability to utilize citrate (Paille et al. 1987).

There is also a high degree of genetic relatedness (moles % guanine plus cytosine and DNA-DNA homology) between environmental isolates and clinical strains of K_. pneumoniae (Woodward et al. 1979; Seidler et al. 1975). The onlyobservable difference is that there is more variability in the genetic values generated by the environmental strains (Seidler et al. 1975).

One concern is that pathogenic K_. pneumoniae can apparently survive for some time in the environment (Knittel et a l . 1977). Another complicating factor is the ability ofjC. pneumoniae and E. col i to exchange resistance factors or R-factors (Allen et a l . 1988; Talbot et al. 1980; Salzman et

al. 1967; Salzman ft Klemm 1966). Klebsiel la pneumoniae also appears to be capable of becoming more antibiotic resistant than E^ coli (Knittel 1975; Salzman et al. 1967). Theability of K_. pneumoniae and E. col i to transfer antibioticresistance in the environment while possible, however, is considered unlikely (Talbot et al. 1980). Klebsiella sp. isolated from the environment tend to show little MAR (Paille et al. 1987; Matsen et al. 1974; Brown ft Seidler 1973).

Klebsiella pneumoniae is considered to be a minor but opportunistic pathogen, with most cases reported beinghospital related and mainly involving patients who are immuno-compromised (Van Kregten et al. 1984; Matsen et al.1974; Eickhoff 1972; Martin et al. 1971; Montgomerie et al.1970). Medical complications due to K_. pneumoniae include urinary and respiratory tract infections (Martin et al. 1971; Montgomerie et al. 1970), septicemia (Davis ft Matsen 1974)and tropical sprue (Klipstein et al. 1977). Cases aregenerally nosocomial and are responsible for a number ofdeaths each year (Brown ft Seidler 1973; Salzman et al. 1967). When isolated from infected patients, the K. pneumoniae isolates are found to be toxin producers with high MAR (Milch et al. 1987; Klipstein et al. 1983; Davis ft Matsen 1974).

Klebsiella pneumoniae has been implicated in causing food borne infections, but here again these were mainlyhospital related (Kiddy et al. 1987; Montgomerie et al.1970). K. pneumoniae has, however, been isolated from foods

not related to hospitals, such as dairy products, vegetables, and meat products (Kaur et al. 1987; Danielsson et al. 1979; Schiemann 1976).

There is no direct evidence to indicate that water borne IC. pneumoniae have caused disease in humans (Duncan ft Razzel 1972; Eickhoff 1972). There is also no evidence that the consumption of raw oysters, even with high numbers of fecal coliforms, has caused K. pneumoniae infections (Paille et al. 1987; Bryan 1980; Brown ft Dorn 1977; Jensen 1956). Klebsiella sp. isolated from summer harvested Louisiana oysters were tested and shown not to produce a significant amount of toxin (Boutin et al. 1986). These investigatorsconcluded that the mere presence of the Klebsiella sp. should be of little concern with regards to being a health hazard to humans, but that total numbers might be.

MATERIALS AND METHODS

Organisms

Environmental strains (17) were isolated from summer harvested Eastern oysters, Crassostrea virginica (Gmelin), commercially harvested from oyster beds in Terrebonne Parish, LA. Clinical strains (7) of jC. pneumoniae were obtained from the Centers for Disease Control (CDC), Atlanta, GA and from the Food and Drug Administration (FDA), Cincinnati, OH. Strains of Klebsiella planticola and Klebsiella terrigena were also obtained from the FDA. Defined strains (2) of K. pneumoniae were obtained from the American Type Culture Collection (ATCC), Rockville, MD. E_. coli controls included an ATCC strain and another defined strain obtained locally. See Table 1 for a listing of organisms used.

Isolation and Testing of Isolates

Shellstock oysters were collected from oyster boats at dockside. Twelve oysters from several sacks were taken, placed in plastic bags, packed in ice, and transported to Louisiana State University, Baton Rouge, LA for testing. At the laboratory, the oysters were cleaned and shucked, as per American Public Health Association (APHA) recommendations (APHA 1984), and then homogenized in sterile Waring Blender

29

30

Table 1: Organisms Used in This Investigation.

Defined Strains Clinical Isolates Oyster Isolates

Klebsiella Klebsiella Klebsiellapneumoniae pneumon iae sp.ATCC 13882 TS9-19’ 31 FLAATCC 13883 788-78* 58 FLA

4210-83 068417Klebsiella 4531-84 068421plant icola 4533-84 068437

4588-84 0684424245-72 4609-84 068449

laKlebsiella 088444terrigena 088454

0884569015-82 088460

56 ERFAEscherich ia 59 ERFA

col i 66 ERFA67 ERFA

ATCC 25922 K- 12

------- 1— m - --- r ; ------- ~T7Z----- -- S rr ,____ n ...

68 ERFA

1 Toxin Producer 3 Type 9

jars (Eberbach Corp., Ann Arbor, MI). From this homogenate, 11 gm were added to sterile 99 ml dilution blanks (0.1* peptone, 0.85* NaCl) to produce a 1/10 dilution. Further dilutions were made, usually 10'3 to 10'3 , to allow for end point determination. Diluted samples were then inoculated into lauryl tryptose (LST) broth (Difco Laboratories, Detroit, MI) to give a 5 tube MPN and incubated at 35°C. After 24 hours, organisms from positive tubes (growth with gas) were transferred to tubes with EC medium (Difco) using sterile applicator sticks (Hastback 1981). Inoculated ECtubes were incubated in coliform incubator baths (Precision Scientific Group, Chicago, IL) set at 44.5°C for 24 hours. Growth from positive EC tubes was streaked for isolation on Levine eosine methylene blue (L-EMB) agar (Difco). Forty- eight hour positive LST tubes were inoculated into EC medium tubes and treated as above. After 24 hours any growth from additional positive EC tubes were again streaked to L-EMB. Levine eosine methylene blue agar plates were incubated at 35°C for 24 hours and then observed for colony types and morphologies. Colonies giving typical Klebsiella morphology (medium to large pink or dark centered mucoid colonies) were picked, streaked on nutrient agar (NA, Difco), and incubated for 24 hours at 35°C. Isolated colony growth streaked on NA slants was loop transferred and suspended in 5 ml of sterile 0.85* NaCl to give a turbid solution. This in turn was inoculated into API 20E biochemical test strips (Analytab

32Products, Plainview, NY) according to manufacturer’s procedures and incubated at 35°C for 24 hours. After the incubation period, the strips were developed and read.Identification was made using the manufacturer’s codingprocedure (Figure 1). Tests for biochemical reactionsincluded ortho-nitropheny1-B-d-galactopyranoside (ONPG) hydrolysis, arginine dihydrolase (ADH) production, lysine decarboxylase (LDC) production, ornithine decarboxylase (ODC) production, citrate (CIT) utilization, hydrogen sulfide (HsS) production, urease (liRE) production, tryptophane deaminase (TDA) production, indole (IND) production, Voges-Proskaur (VP) reaction, gelatin (GEL) 1iquification, glucose (GLU) fermentation, mannitol (MAN) fermentation, inositol (INO)fermentation, sorbitol (SOR) fermentation, rhamnose (RHA)fermentation, sucrose (SAC) fermentation, melibiose (MEL)fermentation, amygdalin (AMY) fermentation, (1+)arabinose (ARA) fermentation, and oxidase (0X1) reaction. Oxidase reactions were performed using Oxidase identification sticks (Oxoid Ltd., Hampshire, England). Oyster isolates used in this study were fecal coliform positive, were urease negative (at 24 hours), and were identified as l(. pneumoniae, using the API system.

In addition to the above, all test organisms were tested for B-glucuronidase production. 4-Methylumbelliferyl-B-d- glucuronide (MUG, Hach Co., Loveland, CO) was added to both LST and EC medium at 100 ug/ml of prepared media. Isolates

33

f l t f i n n c t N u m b e r a s m<ipi 20ESys,em @ s , » d T c r

P l t n n l . .

P h y s ic ia n . U S l D e p t /S e r v ic e .

O N P G A D H LDC 1 2 4

O D C C JT H ,S t 2 4

U R E T O A IN D 1 2 4

V P1

5 h

2 4 h - t - — Hh — — ------ ------+ • —

4 8 h

P ro fileN u m b e r 5 \ H

G E L G L U 2 4

M A N IN O S O n 1 2 4

R H A S A C M E L 1 2 4

A M Y A B A O X I 1 2 4

------ -t- + -— •f' -f- — - ------- -f' —

U E 5* [5 s]N O , G A S M O T M A C O F -O O F -F

2 4 h

4 8 h

A d d i t io n a l In fo rm a t io n

IfiiViC +-t---I d e n t if ic a t io n

^ • go// id.

cipi20E'System

R e fe re n c e N u m b e r Z M 2 S , .. 4rcc.

P a tie n t . D a ta 9- ?- & ?P h y s ic ia n . D e p t./S e rv ic e .

O N P G1

A D H2

LO C4

O D C1

C IT2

H ,S4

U R E1

T D A2

IND4 VP1

G E L2

G L U4

M A N1

IN O2 son4

R H A1

SA C2

M E L4

AM Y1

A R A2

0 X 14

5 h

2 4 h -- + — + -- H h — -- — -t- -h + ■t + " 1 “ ■h + -4 8 h

P rofileN u m b e r 5 a . 1 5 7 - A

N O , G A S M O T M A C O F -O O F -F

2 4 h

4 8 h

□

A d d it io n a l In fo rm a t io n

IH4 V i C . + 7~

Id e n tif ic a t io n

K • f n e u r o n e x c e . / f& j- f - i<L

Figure 1: API 20E Test Strip Identification Codes.

were loop inoculated into LST and incubated at 35CC. At 24 hours, results were determined by looking for normal LST positive reactions and for fluorescence (evidence of B- glucuronidase production) under UV light (Gelman Instrument Co., Ann Arbor, MI) at 375 nm. Organisms from all LST tubes showing growth, regardless of the coliform and MUG reactions, were inoculated into EC medium and incubated in coliform incubator baths at 44.5°C. Results were read at 24 hours by looking for normal EC medium positive reactions and again by observing for fluorescence.

A second commercial preparation, Minimal Media ONPG-MUG (MMO-MUG) or Colilert (Access Analytical Systems, Branford, CT), also, was used. This product comes in ready-to-use sterile tubes designed to test water samples. For the purposes of this investigation, isolates were suspended in 10 ml of sterile 0.8536 NaCl to make turbid solutions. These solutions were then inoculated into the Colilert tubes and incubated at 35°C for 24 hours as per manufacturer's instructions. Since these tubes have no durham tubes to detect gas production, presence of fecal coliforms is determined by ONPG hydrolysis by B-galactosidase. Fecalcoliform positive tubes develop a yellow color. Production of B-glucuronidase was again determined by positive fluorescence.

Deoxyribonucleic Acid (DNA) Isolation35

A modified version of Maumer’s (1961) DNA extraction procedure was used. Cultures grown for 24 hours on brain- heart infusion (BHI) agar (Difco) slants were loop inoculated into 100 ml of BHI broth. After a further 24 hours of growth, 2 ml of culture were added to each of eight 500 mlRoux flasks containing 60 ml of BHI agar. Roux flasks wereincubated at 35°C overnight (about 18 hours). Twenty milliliters of filter sterilized saline-ethylenediaminetetraacetate (EDTA, SE) with 0.1% protease (Sigma Chemical Co., St. Louis, M0) were then added andincubated for 1 hour at 35°C. The composition of thesolutions used in the DNA extraction procedure is listed in Table 2. Sodium azide (0.2%) was added to some of the solutions as a preservative.

The bacterial suspensions were collected (poured) from the Roux flasks, 50 ml of isopropanol added, and the suspensions agitated for about 10 minutes. Suspensions were centrifuged in 250 ml polycarbonate centrifuge bottles (Nalgene Co., Rochester, NY) at 8000 RPM, using a GSA rotor (10,400 relative centrifugal force, RCF), in a Sorval RC2C low temperature centrifuge (Du Pont Co., Newtown, CT), at 0°C for 15 minutes. The supernatants were discarded and thepellets resuspended in 100 ml of SE. These suspensions werecentrifuged again as above, except the pellets were

36

Table 2: DNA Solutions Used in This Study1

Solutions Composition

SE 58.4 gm NaCl (0.5M), 58.5 gm EDTA (0.1M), 4.0 gm NaN3 (0.296), 2000 ml DH20, pH 8.0 (w/10N NaOH)

NP 175.6 gm NaClOa (5M)CIA 480 ml Chloroform, 80 ml Iso Amyl Alcohol

(24:1)CSC 8.76 gm NaCl (1.5M), 4.4 gm Tri Sodium

Citrate (TSC) (0.15M), 0.2 gm NaN3 , 100 ml DH20, pH 7.0

SSC 10 ml CSC, 90 ml DHZ 0DSC 5 ml CSC, 495 ml DHZ 0Acetate-EDTA 24.6 gm Na Acetate (3M), 0.03 gm EDTA,

0.2 gm NaN3 , 100 DHr 00. 1% Protease 0.1 gm Protease (Sigma, P-4880), 100 ml

SE (filter sterilize)0.5% Protease 0.025 gm Protease, 5 ml DHZ03.096 Lysozyme 0.15 gm Lysozyme (Sigma, L-6876), 5 ml

DHZ 0

in•o Lysozyme 0.025 gm Lysozyme, 5 ml DH200.496 RNase 0.01 ml Rnase (Sigma, R-5000), 0.04 gm

NaCl, 5 ml DH20 (DNase free by heat inactivation at 80°C for 10 minutes)

1 From Maumer 1961

resuspended in 50 ml of SE containing 2556 sucrose.To each suspension, 2 ml of 3% lysozyme (Sigma) were

added and the mixtures incubated at 35°C with shaking at 200 RPM on a Gyrotory Shaker (New Brunswick Scientific Co., Inc., New Brunswick, NJ) for 30 minutes. Ten milliliters of 2596 sodium dodecyl sulfate (Sigma) were then added and the mixtures incubated for 30 to 60 minutes at 60°C with occasional mixing. The reaction mixtures were then cooled to room temperature (20° to 25°C). Subsequently, 12.5 ml of 5 M sodium perchlorate (NP) (Fisher Scientific, Pittsburgh, PA) were added and the solutions gently mixed. Sixty milliliters of chloroform-iso-amyl alcohol (CIA) (24:1) were then added, the solutions gently swirled to mix, and stored overnight at 4° C.

Suspensions were gently swirled to mix the aqueous and organic phases, and then centrifuged in 250 ml plastic bottles using a GSA rotor at 10,000 RPM (16,300 RCF) for 20 minutes at 0°C. Using wide mouth pipettes the upper (aqueous) phases was removed to 250 ml screw top flasks. Then, 10 ml of NP were added, the suspensions mixed, and then 60 ml of CIA added. Reaction mixtures were mixed again and centrifuged in 150 ml Corex bottles (Corning Glass Works, Science Products, Corning, NY) using a GSA rotor at 5000 RPM (4,080 RCF). This last step was repeated until theprecipitation at the interface of the phases had disappeared and the upper phase were clear. The upper phases were then

transferred to 250 ml graduated cylinders and two volumes of cold (-20°C) 9596 ethanol (ETOH) layered on top. Using longglass rods, the crude deoxyribonucleic acid (DNA) from each cylinder was spooled out by gently swirling the glass rod at the interface between the DNA solution and the cold 9596 ETOH. When the reaction solutions became uniform (no interface) and it appeared that no more DNA was being spooled, the DNA was gently pressed out to remove excess fluid by gently rolling it on the inside of the graduate cylinder above the spent solution. The spooled DNA was rinsed with cold 9596 ETOH and allowed to air dry at room temperature. The DNA was then resuspended in 8 ml of standard saline citrate (SSC, 1/10 concentrated saline citrate, CSC) and incubated overnight at 4° C .

The following day, DNA suspensions were warmed to room temperature. If the DNA suspensions had not completely resuspended, they were gently swirled until resuspension occurred. After this, 0.496 RNase (Sigma) was added to the suspensions, which were then mixed, and incubated at 35°C for one hour with shaking, at 100 RPM. To the reaction mixtures 0.596 lysozyme was added, mixed, and incubated as above for an additional half hour. Next, 0.596 protease was added to the mixtures, mixed, and incubated as above. Finally, 1.6 ml of NP then 10 ml CIA were added to the mixtures which were gently swirled, and then incubated for 1 hour at 4°C. After this incubation period, the mixtures were centrifuged in 30

ml Corex tubes (Corning) using an SS-34 rotor at 10,000 RPM (12,000 RCF). The DNA was collected by spooling, as before, resuspended in 9 ml of dilute saline citrate (DSC, 1/10 SSC), and stored at 4°C overnight. The next day, the suspensions were warmed to room temperature. Once again, if the DNA suspensions were not completely dissolved, they were gently mixed until they were resuspended. One milliliter of acetate-EDTA was then added and the mixtures were incubated for two hours at 4°C. The DNA mixtures were placed in 100 ml graduated cylinders and two volumes of cold (-20oC) isopropanol layered on top. The DNA was then spooled out as before. If the alcohol solutions were cloudy after spooling the DNA, the DNA was resuspended in DSC as above, and the spooling procedure repeated until the alcohol solution became clear. Finally the DNA was washed free of acetate with a series of ETOH washes, 70%, 80%, and 95%, and resuspended in 5 ml of DSC. The DNA mixtures were stored at -85°C in a Revco ultra low freezer (Rheem Manufacturing Co., Ashville, NC) until DNA analysis procedures were performed.

Moles % Guanine Plus Cytosine (Mol% G+C)

The moles % guanosine plus cytosine (mol% G+C) content was determined by thermal denaturation, using a Gilford model 2400 spectrophotometer with a model 2527 thermoprogrammer (Ciba-Corning Diagnostics Corp., Gilford Systems, Oberlin,

OH). Prepared DNA samples were first diluted with DSC to an absorbance of 0.600 at 260 nm. Samples (0.32 ml) were loaded into cuvettes, placed into the thermoset, heated to a predenaturation temperature cf 60°C, and then heated at a controlled rate of 1°C per minute until denaturation was completed (approximately 90°C). Data were collected using the instrument's mechanical plotter. Median melting temperatures (TM ) were determined by identifying the inflection point of the melting curves (Figure 2). The temperature values at the T* were used in the calculation of mol% G+C content. The formula used was that of Norgad & Bartel1 (1978):

Mol% G+C = [(Tm /50.2) - 0.99) x 100

DNA samples were tested in triplicate and the mean T„ used tocalculate mol% G+C contents.

A repeat run was performed using a Gilford Response II spectrophotometer (Ciba-Corning). DNA samples werestandardized to an absorbance of 1.000 at 260 nm, as opposed to the 0.600 used above. This instrument stored data in its internal memory. The T* values were calculated by the instrument as the first derivative of the melt curves (Figure3 and 4). The mol% G+C were calculated as above.

40 30 20SO 1000 70100

40 30 2000 1060 7000-

-)►30 20-50 4070- 001 0 0 SO 80

3040 20S O6010O S O 00

SO- 40- 30 201 0 0 SO fa eo-80

M l

Relative Absorbance (260 rm)

Figure 2: Mol% G+C Oenaturation Plot from a Gilford 2400 Spectrophotometer,

Taiperatu

re (°C

)

421 . 501 . 4 0

K. pneuTpntaeE. colt

- 1.1088 .065 .0 T E M P E R A T U R E (°c)

Figure 3: Mol1* G+C Denaturation Plot from a Gilford Response II Spectrophotometer.

0 . 0 6 -

- 0 . 0 1- 0.0188.065 . 0 T E M P E R A T U R E <°c)

Figure 4: First Derivative Plot from a Gilford Response I I Spectrophotometer.

43DNA-DNA Homology

Homology determinations were accomplished using thethermal renaturation procedure of Norgad & Bartell (1978), with a Gilford model 2400 spectrophotometer. Chromosomal DNA samples, standardized to an absorbance of 3.000 at 260 nm, were first fragmented to a uniform size of about5.0 x 10* daltons by passing each DNA sample individuallythrough 25 gauge needles lOx against the side of beakers, using maximum hand pressure. Fragmented, standardized DNA was then diluted 1:1 in 12x SSC containing 50% spectro grade formamide (Eastman Kodak Co., Rochester, NY). One sample was compared to the reference (clinical isolate 4210-83) in each run. DNA from the reference and sample were compared to a 1:1 mixture of the reference and sample. DNA mixtures (0.32 ml) were pipetted into cuvettes, placed in the thermoset, and heated to 95°C. After holding the DNA at this denaturation temperature for 5 minutes, the samples were cooled as rapidly as possible (about two minutes) to the renaturation temperature (T0r ) of 60°C. At this point, the absorbance profile, at 270 nm, was monitored for 10 minutes. Data were collected using the instrument’s mechanical plotter. The slopes of the linear portion of the renaturation curves were used to determine % homology by comparing the 1:1 DNA mixture to the reference strain and the DNA sample being tested (Figure 5). Calculations of % homology were made

44

30 207 0100

' U t t

T>4-

10b 30 10S O 2060eo 70

3*SO 10so■1C BO 00

1040 30 2070- SOIOO 00 008 0

Relative Absorbance (270 nh)

Figure 5: DNA Homology Plot from a Gilford 2400 Spectrophotometer.

Terj>e

rature

(°C

)

using the following formula (De Lay et al. 1970):45

% Homology = L4V„1X - (V, + V2 )/2(V,V* )' 1 '3 ] 100where: V, = slope DNA 1

V2 = slope DNA 2V«ix = slope of mixture of DNA*s 1 and 2

As with the mol% G+C tests, DNA samples were tested in triplicate and the mean of the slopes used in the above formula.

A second run was performed using a Gilford Response II spectrophotometer. This instrument allowed the testing of two samples of DNA to the reference C1C. pneumoniae, ATCC 13882) at one time and determined the slopes as the change of absorbance per minute (dA/min) (Figure 6). Slopes were used as above to calculate % homologies.

A flow chart of the entire DNA analytical procedure canbe seen in Figure 7.

46

S A M P L E S =

LI N E A R FIT

RATE d A/M IN - 0 . 0 1 1 8 - 0 . 0 1 1 9 -0.0106 -0.0061STD. DEV. 0 . 0 0 0 6 0. 0 0 0 6 0.0008 0.0003UNITS. FCT 0 . 0 1 1 8 0 . 0 1 1 9 0.01O6 0.0061LI N E A R TO i ’4 YES YES YES YES

N O N - L I N E A R FIT

IN1T RATE STD. DEV. UNITS. FCT

Figure 6: dA/Minute Determinations by a Gilford Response II Spectrophotometer.1 - type strain DNA2 - sample 1 DNA3 - sample 2 DNA4 - type x sample 1 (1:1)5 - type x sample 2 (1:1)

5

0.0085 0.0004 0.0085YES

Day 1: Inoculate LST with Oyster Sa»7iplesII

Day 2: Inoculate EC with LST PositivesII

Day 3: Streak EMB with EC PositivesII

Day 4: Select Colonies & Inoculate BHI SlantsII

Day 5: Inoculate BHI Broth I I

Day 6: Inoculate Roux FlasksII

Day 7: Collect & Lyse Cells I I

Day 8: Collect Crude DNA I I

Day 9: Clean up & Concentrate DNAII

Day 10: Final Clean up & Preparation of DNA ) iI t

Moles % G+C DNA-DNAHybridization

Figure 7: Flow Chart for DNA Analysis.

RESULTS AND DISCUSSION

Biochemical reactions on 6 defined strains, 7 clinical isolates and 17 oyster isolates, determined by API 20E test strips, are presented in Table 3. Klebsiella planticola and K. terrigena were not identified as such using the API 20Ebiochemical test strip identification index. Oyster isolates selected for this study were all urease negative at 24 hours, but 9 of 17, or 53%, tested positive after 48 hours. Paille et al. (1987) indicated that 69% of Klebsiella sp. isolatesfrom their work were urease negative. Nonclinical, urease negative K. pneumoniae have been previously identified,predominantly from vegetable and water samples (Naemura et al. 1979; Bagley & Seidler 1977). The jC. planticola and K. terrigena isolates were previously characterized as urease positive (Bagley et al. 1981; Izard et al. 1981). Thispresent study, however, differed in that l(. terrigena was found to be urease negative.

Previous work on antibiotic resistance on some of the oyster and clinical isolates tested (unpublished data) is presented in Table 4. Antibiotic resistance was determined using the procedure of Bauer et al. (1966). Two of theclinical isolates had MAR values of 6 and 10 (of 18antibiotics tested) while most had MAR values of 3 or 4. All of the oyster isolates tested had antibiotics tested) while most had MAR values of 3 or 4. All of the oyster isolates

48

49

Table 3: API 20E Test Results of This Investigation.API 20E Biochemical Reactions2

Organisms ONPG ADH LDC ODC CIT H2S URE TDA IND VP GELE. coli25922 + - +K-12 + - +

K. planticola4245-72 + + +

K. terrigena9015-82 + + +

K. pneumoniae13882 + - +13883 + - +

Clinical IsolatesTS9-19 + - +788-78 + - +4210-83 + - +4531-84 + - +4533-84 + - +•4588-84 + - +4609-84 + - +

Oyster Isolates31 FLA + - +58 FLA + - +068417 + - +068421 + - -068437 + - +068442 + - +068449 + - +

la + - +088444 + - +088454 + - +088456 + - +088460 + - +56 ERFA + - +59 ERFA + - +66 ERFA + - +67 ERFA + - ♦68 ERFA + —

++

++

++

Urease Positive at 48 hours.See p. 31 for Explanation of Abbreviations.

50

Table 3: API 20E Test Results (Cont.).API 20E Biochemical Reactions

Organisms GLU MAN INO SOR RHA SAC MEL AMY ARA OX IE. coli25922 + + - + + -K-12 + + + - - - -

K. planticola4245-72 + + + + + + + + + -

K. terrigena9015-82 + + + + + + + + + -

K. pneumoniae13882 + + + + + + + + + -13883 + + + + + + + + + -

Clinical IsolatesTS9-19 + + + + + + + + + -788-78 + + + + + + + + + -4210-83 + + + + + + + + + -4531-84 + + + + + + + + + -4533-84 + + + t + + + + + -4588-84 + + + + + + + + + -4609-84 + + + + + + + + + -

Oyster Isolates31 FLA + + + + + + + + + -58 FLA + + + + + + + + + -068417 + + + + + + + + + -068421 + + + + + + + + + -068437 + + + + + + + + + -068442 + + + + + + + + + -068449 + + + + + + + + + -

la + + + + + + + + + -088444 + + + + + + + + + -088454 + + + + + + + + + -088456 + + + + + + + + + -088460 + + + + + + + + + -

56 ERFA + + + + + + *■ + + -59 ERFA + + + + + + + + + -66 ERFA + + + + + + + + + -67 ERFA + + + + + + + + + -68 ERFA + + + + + + + + + -See p . 3 1 for Explanation of Abbreviations.

51

Table 4: MAR of Selected Organisms Used in This Study.Clinical Isolates if

Antibiotics 4210 4531 4533 4588 4609ft

Resist.Nitrofurazone (100 ug) + ' + + + + 0Kanamycin (30 ug) + + + - + 1Neomycin (30 ug) + + + + + 0Polymyxin B (50 units) + + + + + 0Oxytetracycline (30 ug) + + + - + 1Chlortetracycline (30 ug) + + + + + 0Nitrofurantoin (100 ug) + + + + + 0Vancomycin (30 ug) - - - - 5Cephalothin (30 ug) + + + + + 0Streptomycin (10 ug) - + + + + 1Sulfathiazole (1.0 mg) - + - + 3Chloramphenicol (30 ug) + + + - + 1Carbenicil1 in (50 ug) - - + - + 3Erythromycin (15 ug) + + - - 3Ampicillin (10 ug) - - + - - 4Nalidixic Acid (30 ug) - + + + + 1Tetracycline (30 ug) + + + - + 1Gentamicin (10 ug) + + + — + 1MAR Ratio

. ii >■ ,, ,, , .6/ 18 3/18 3/ 18 10/ 18 3/18 12/ 18

+ = Susceptible - = Resistant

52

Table 4: MAR of Selected Organisms (Cont.)Oyster Isolates

06 06 06 06 06 #Antibiotics 8417 8421 8437 8442 8449 Resist.Nitrofurazone (100 ug) . i + + + + 0Kanamycin (30 ug) + + + + + 0Neomycin (30 ug) + + + + + 0Polymyxin B (50 units) + + + + + 0Oxytetracycline (30 ug) + + + + + 0Chlortetracycline (30 ug) + + + + + 0Nitrofurantoin (100 ug) + + + + + 0Vancomycin (30 ug) - - - 5Cephalothin (30 ug) + + + + + 0Streptomycin (10 ug) + + + + + 0Sulfathiazole (1.0 mg) + + + + 0Chloramphenicol (30 ug) + + + + + 0Carbenicillin (50 ug) - - - - - 5Erythromycin (15 ug) - + + + - 2Ampicillin (10 ug) - - - - - 5Nalidixic Acid (30 ug) + + + + + 0Tetracycline (30 ug) + + + + + 0Gentamicin (10 ug) + + + + + 0MAR Ratio 4/18 3/ 18 3/ 18 3/18 4/ 18 4/18

' + = Susceptible- = Resistant

tested had MAR values in the 3 to 4 range. Low MAR in environmental isolates of Klebsiella have been documented by other studies (Matsen et al. 1974; Brown ft Seidler 1973).The data indicate that the Klebsiella sp. from thispresent investigation were universally resistant toampicillin, carbenicillin and vancomycin. Strains of 1C. pneumoniae have been shown to be resistant to ampicillin and vancomycin in other studies (Bagley et al. 1981; Talbot et al. 1980; Matsen et al. 1974). Since pathogenic strains of 1C. pneumoniae with high MAR have been shown to be able to exist for long periods in the environment (Talbot et al. 1980; Knittel et al. 1977), it could be expected that if theKlebsiella sp. isolated from oysters were from recent fecal contamination that at least some of the MAR values should have been higher. Low MAR do not necessarily indicate that sewage is not present but high MAR would indicate the presence of human sewage or contamination from animals treated with antibiotics. Another aspect is that JC. pneumoniae and E_. coli have the ability to exchange R-factors (Allen et al. 1988; Talbot et al. 1980; Salzman et al. 1967;Salzman et al. 1966). However, the ability of these organisms to exchange R-factors in the environment, while possible, is considered unlikely (Talbot et al. 1980).Escherichia coli associated with enteric disease has been shown to have high MAR. It has been proposed that MAR be used to screen foods when high-risk sources of fecal

contamination is suspected (Krumperman 1983). This argument would probably also hold for Klebsiella sp. isolated from oysters.

All the oyster isolates chosen for the DNA analysis were biochemically identified as l£. pneumoniae, and were fecal coliform positive. Cultures were maintained on BHI agar at room temperature (20° to 25°C) and retested after several passages. Results indicated that 65% of the organisms lostthe ability to produce acid and gas from lactose at 44.5°C (Table 5). The significance of this is not clear but may suggest adaptation or mutation. Loss of the ability toproduce a positive fecal coliform reaction may reflect whatnaturally occurs to Klebsiella sp. held in the environment for long periods of time at low temperatures. In addition, it may be speculated that high temperatures in theenvironment may select for or induce fecal coliform negative Klebsiella sp. to become fecal coliform positive. This may account for the increased presence of Klebsiella sp. during the warmer months and its reduced presence during the cooler months.

The isolates were also tested for their MUG reaction. In all cases the clinical and oyster isolates showed negative fluorescence (Table 5). Reactions to MUG were similar inboth LST and EC medium. MMO-MUG appeared to be lesssensitive in its ability to detect E. col i K-12. Thereaction for E. coli K-12 required 48 hours to show positive

55

Table 5: Fecal Coliform and MUG Reactions from This Study.Fecal Coliform Reactions

Organisms Preliminary Retest MUGE. coli25922 + + +K-12 + + +(slow)

K. planticola 4245-72

1C. terr igena9015-82 -

K. pneumoniae13882 + +13883 -

Clinical IsolatesTS9-19 + +788-78 -4210-83 -4531-84 -4533-84 -4588-84 +4609-84 + + -

56

Table 5: Fecal Coliform and MUG Reactions (Cont.)Fecal Coliform Reactions

Organisms Preliminary Retest MUGOyster Isolates

31 FLA58 FLA 068417 068421 068437 068442 068449

la 088444 088454 088456 088460 56 ERFA59 ERFA66 ERFA67 ERFA68 ERFA

+

+

Ratio 17/17 6/17 0/ 17

fluorescence. These results simply confirm what has already been shown in other studies (Perez et al. 1986; Feng &Hartman 1982; Kilian & Bulow 1976) and serve to reinforce that MUG would more than likely not identify j<. pneumoniae if incorporated into the fecal coliform test. One study thatdid recover Klebsiella from a positive tube was unable to confirm that it was responsible for the positive reaction when retested (Koburger a Miller 1985). Those authors suggested a possible synergistic interaction reaction had occurred. Considerations not mentioned include thepossibility that the organism actually responsible for the positive reaction was not recovered or the possibility of autofluorescence due to dirty glassware.

Concerns with MUG include questions about falsepositives due to other enterobacteriaceae, includingSalmonella, Shigella, and Citrobacter (Perez et al. 1986),and false negatives, as compared to the standard fecalcoliform test. Since the false positives are associated with known pathogenic organisms, their detection, if they arepresent, should not cause major concern. At the temperature used to test shellfish for fecal coliforms, 44.5°C, the false positive rate of the standard fecal coliform test in one study was as high as 4096 (Fishbein & Surkiewicz 1964). Of more concern is the problem of false negatives and the possibility of not detecting the true levels of E. colipresent. In general the false negative rate is <1096 (Iritani

& Inzana 1988; Andrews et al. 1987; Rippey et al. 1987; Perez et al. 1986; Robinson 1984; Feng & Hartman 1982; Kilian &Bulow 1976). This value must be compared with the falsenegative rate of the standard test, which also has been questioned for underestimating actual fecal coliform levels (Poelma et al. 1988, Cook & Pabst 1984). Since using MUG is credited with increasing the sensitivity of the fecal coliform test by detecting E_. col i in conditions where thepresence of E. coli is masked by the growth of otherorganisms, the use of MUG may result in a more accurate reporting of the levels of fecal contamination (Rippey et al. 1987; Moberg et al. 1985). Also, one study indicated thatMUG was 100% effective in detecting enteropathogenic E. coli(Feng & Hartman (1982). The failure of MUG to detect all E_.coli present in a sample may be due to environmental strains. Environmental strains of E_. col i should not be of concern in predicting the sanitary quality of samples being tested.

DNA work was carried out to determine the relatedness of the clinical and oyster isolates compared with standards. Two runs were undertaken. In each, samples were tested in triplicate and the mean of the three tests used to estimate the parameters of interest. The second run was undertaken to strengthen the data, because of uncertainties encountered during the first run. In the first run, the use of a mechanical plotter was suspected of being a source of unintentional investigator bias, due to subjective analysis

59of the data and mechanical and/or electrical problems. In the second run, a different instrument was used, one that stored data points in internal memory and plotted and calculated parameters using microprocessors, thus reducing the chance for any investigator-introduced bias.

Preliminary analysis of the purified DNA samples indicated relatively high DNA concentrations with some extraneous contaminants, possibly due to polysaccharides and protein not removed by the purification procedure. The mean of the Az6 o /Aa ao ratios averaged 1.8, with the mean of the protein levels averaging <1 ug/ml.

The median melting points (T„), calculated mol% G+C, andsimple statistics are presented in Tables 6, 7, and 8. Ingeneral, the values generated in the second run were slightly higher than the first but were more consistent than findings of the first run. The two runs also differ in that thesecond more clearly differentiates K. plant icola and 1C. terrigena from the oyster isolates, making it appear that the oyster isolates are more closely related to the clinical isolates or K. pneumoniae.

The data indicate that the defined, clinical, and oyster isolates mol% G+C values generated are higher than thereported values for K_. pneumoniae, 52% and 56% (Cowan 1974). The values generated by this present investigation may reflect an differences due to the formula used to calculate the mol% G+C, since the T„ is dependent on the cation

60

Table 6: Moles % Guanine Plus Cytosine of the Strains Tested.

OrganismsI*1 Run 2" " Run

T„ G+C Th G+CE. coli25922 76.2 52.8K- 12 76.3 53.0 76.2 52.8

K. planticola4245-72 78.3 57.0 78.6 57.6

K. terrigena9015-82 79.0 58.4 79.6 59.6

K. pneumoniae13882 — 79.2 58.313883 78.8 58.0

Clinical IsolatesTS9-19 78.7 57.8 79.2 58.8788-78 78.4 57. 1 79.3 59.04210-83 78.9 58. 1 79.3 59.04531-84 78.6 57.6 79.3 59.04533-84 78.6 57.6 79.4 59.24588-84 77.9 56. 1 79.3 59.04609-84 78.6 57.6 79.6 59.6

Oyster Isolates31 FLA 78.7 57. 8 79.4 59.258 FLA 79.0 58.4 79.3 59.0068417 78.6 57. 6 79.5 59.4068421 78.0 56.3 79.4 59.2068437 78.7 57.7 79.2 58.8068442 77.9 56.9 78.9 58.2068449 79.2 58.7 79.2 58.8

la 79.2 58.7 79.3 59.0088444 78.3 57.0 79.3 59.0088454 78.4 57.2 79.4 59.2088456 78.4 57.2 79.0 58.4088460 78.6 57.6 79. 1 58.656 ERFA 78.6 57.6 79.2 58.859 ERFA 78.6 57.6 79. 1 58.666 ERFA 78.4 57.2 79.3 59.067 ERFA 77.9 56.2 79.3 59.068 ERFA 78.5 57. 4 79.4 59.2

61

Table 7: The Mol% G+C of Clinical Isolates Tested.1* 1 Run 2" ° Run

I tern T„ G+C Th G+C

MeanSD1Range

78.50.29

77.9-78.957.4 0.60

56.1-58.179.3 0. 12

79.2-79.659.1 0.24

58.8-59.6' Standard Deviation

Table 8: The Mol% G+C of Oyster Isolates Tested. F-* Run 2"d Run

Item T* G+C T» G+C

Mean 78.5 57.4 79.3 58.9SD 0.37 0.73 0.15 0.31Range 77.9-79.2 56.2-58.7 78.9-79.5 58.2-59.4

concentration of the buffer in which the DNA is suspended (Gill is et al. 1970; Mandel et al. 1970; Mandel & Maumer I960; Szybalski 1967). In general, the higher the cation concentration the higher the T„ for any given DNA. The values of this study are, however, in line with the findings of other studies, 54% to 59% (Seidler et al. 1975; Gillis et al. 1970). Also, the values for E. coli K-12 wereessentially the same as in other studies (Norgard & Bartell 1978; Mandel et al. 1970).

Strictly based on the mole % G+C data, the onlyconclusion that can be made is that the organisms in question belong to the genus Klebsiella. Species identification can not be inferred, although the two known environmentalisolates seem to be set apart, at least by the second run. These results confirm those of Paille et al. (1987) who reported a mean mol% G+C value of 57.8% for urease negative Klebsiella sp. isolated from summer harvested Louisiana oysters (using a Gilford 2400 spectrophotometer). The mol% G+C value for K_. planticola is higher than the reported value of about 55% (Bagley et al. 1981), but in keeping with their findings, values were lower than those reported for K. pneumoniae.

The DNA-DNA homology results and simple statistics are presented in Tables 9, 10, and 11. A clinical isolate, 4210- 83, was arbitrarily selected based only on it being ureasepositive. For the second run, a defined strain of

63

Table 9: Levels of Homology of Organisms Examined.Percent Homology

Organisms 1* 4 Run1 2" a Run'E. coli25922 13. 1K-12 12.7 15.9

K. planticola4245-72 25.0 41.4

K. terrigena9015-82 13.0 30.8

K. pneumoniae13882 100.013883 80.2

Clinical IsolatesTS9-19 86. 9 78.9788-78 97. 1 73.54210-83 100.0 75.94531-84 81. 5 92.44533-84 25. 6 21.64588-84 87.3 87.34609-84 89. 1 87.6

Oyster Isolates31 FLA 87.2 84.358 FLA 99.7 87.6068417 75.4 86. 1068421 62.5 69.3068437 86.3 90.7068442 94. 1 72.2068449 69.5 77.0

la 63.7 64.7088444 80.0 89.5088454 88.0 94.2088456 86.9 82.5088460 90. 3 88.656 ERFA 84.8 85.759 ERFA 78.4 81 .466 ERFA 71.0 83.267 ERFA 82.2 82.968 ERFA 80.6 76.7

4 Ref. 4210-83

' Ref. 13882

64

Table 10: Level of Homology of Clinical Isolates Tested.Percent Homology

I tern 1** Run’ 2" “ Run2

Mean 88.4 82.6SD 5.0 6.9Range 81.5-97.1 73.5-92.4

' Excludes 4210 & 4533 2 Excludes 4533

Table 11: Level of Homology of Oyster Isolates Tested.Percent Homology

Item 1*T Run 2”a Run

Mean 81.2 82.2SD 10.0 7.7Range 62.5-99.7 64.7-94.2

K. pneumoniae (human isolate), ATCC 13882, was used as the reference. Comparing the reference strains to coli andthe two known environmental isolates showed that different species could easily be differentiated using the renaturation technique used in this study. Also, a test of the relatedness of K. planticola and K_. terrigena , in the first run, showed only 30.1% homology.

Actual numbers varied in the second run, but in general were more consistent than in the first run. Part of the variation may have been due to the use of different reference strains.

Except for 4533-84, the degree of relatedness of the clinical isolates to the reference was >_81.5% for the first run. The odd isolate showed only 25.6% homology to 4210-84, indicating either a different species or a problem with the DNA itself. No attempt was made to confirm the identification of this or any of the other clinical isolates beyond the biochemical tests and DNA analysis. The clinical isolates in the second run, with the exception of one, also showed a high degree of relatedness, >73.5%. Again, 4533-84 showed a low degree of homology, about 21.6%, to the reference. With both runs showing about the same results, they are probably real. At 20% to 25% homology, 4533-84 can not be K. pneumoniae and is probably one of the other Klebsiella species, or possibly Enterobacter aerogenes.

With regards to the oyster isolates, they were related

to 4210-83 at >_62.596 and related to ATCC 13882 at >64.796. Based on percent homology, as it relates to relatedness of species, these numbers are too high to suggest separate species. At best, some of the isolates could representenvironmental subspecies of 1C. pneumoniae. It is probably safe, however, to say that they are the same species.

Test runs comparing ATCC 13883 to 4588-84 (urease positive) and to 31 FLA (urease negative) showed 83.796 and 87.996 homology, respectively. These values vary slightly from those generated by ATCC 13882 but are probably not significantly different.

Compared to the results of studies, by other investigators, comparing environmental water isolates to type strains, the results of this study are comparable, in that all show homologies at or greater than 6396 (Woodward et al. 1979; Seidler et al. 1975). Homology results of 1C. planticola, compared to previously reported values of 796 to 2996 (Bagley et al. 1981), are similar for the first run, 2596 but higher for the second, 4196. Results of the environmental strain 1C_. terrigena showed less homology than a previous study; 1396 and 3196, as opposed to 4896 to 6396 (Izard et al. 1981).

Based strictly on the DNA results (Figures 8 and 9), it must be concluded that the oyster isolates are K. pneumoniae, since all DNA homology values generated were well above the generally accepted 5096 cutoff to consider an organism as

67C B A

I < > I <-> I <--------- > II---- |---|--|---|---|---|---|--- |---1---|0 10 20 30 40 50 60 70 80 90 100

% DNA Homology

A = Same SpeciesB = Same Species or Possible Subspecies C = Closely Related Species

Figure 8: Percent DNA Homology for Taxonomic Groupings (Johnson 1984).

K.terrigena

IE. I 1C. 1C. pneumoniaecoli I planticola 13883 13882* * * # ■* *

- I -----|---- |---- |---- |---- |---- |-----|---- |----|10 20 30 40 50 60 70 80 90 100

% DNA Homology of Type Strains

4533-84 * ** * # * «I---- |--|---|---|---|---|---|--- |---|-- |

0 10 20 30 40 50 60 70 80 90 100% DNA Homology of Clinical Isolates

** ****#

* **** **** *I---- |--|---(---- |--|---|-- |--- |---- |---|0 10 20 30 40 50 60 70 80 90 100

% DNA Homology of Oyster IsolatesFigure 9: DNA Homology Data (Second Runh

belonging to a separate species (Ewing 1986; Johnson 1984; Norgard & Bartell 1978). Whether the oyster isolates are from human sewage, long term fecal survivors, or environmental variants, it is clear that using generally available techniques, they can not be differentiated from clinical isolates except possibly by using MAR.

Based on the data from this study and other investigations, there appears to be reasonable doubt about the significance of the presence of K_. pneumoniae in predicting the legitimate sanitary quality of oysters and other seafoods in general. Also, there appears to be no direct health threat due to these organisms when found in shellfish. Eickhoff (1972), in a review of the Klebsiella problem, could not find evidence for a health hazard due to waterborne organisms. Also, Boutin et al. (1986) showed thattoxin production of Klebsiella sp. isolated from summer harvested oysters was not significant and doubted any health threat due strictly to its presence.

Taking the above into account, along with the apparent ubiquitous presence of K_. pneumon iae in the environment, K_. pneumoniae does not seem to be an appropriate indicator of human sewage. Klebsiella pneumoniae have been found in the absence of pathogens, namely in unpolluted, pristine areas (Rivera et al. 1988; Duncan & Razzel 1972), and it seems to be able to survive for long periods of time in the environment and may actually reproduce there (Lopez-Torres et

al. 1987; Hiraishi et al. 1984; Vasconcellos & Swartz 1976).With this in mind, it can be argued that the fecal

coliform test as it is now used should be modified to distinguish K_. pneumoniae. This can be done by incorporating MUG into the test. To avoid the false positive problems caused by B-glucuronidase naturally present in shellfish tissue, the MUG reagent should be incorporated into the EC medium. This will allow the direct detection of E. coli and differentiation from K_. pneumon iae, which also gives a fecal coliform positive reaction, but which is MUG negative. The importance of this was shown in one study that indicated that 90% of non fecal Klebsiella were incorrectly identified as being fecal coliform positive (Caplenas & Kanarek 1984). A question concerning MUG that must be addressed is the presence of E. coli which would be not of concern in predicting the sanitary quality of shellfish and their environment. These E. col i might represent those that are not detected by MUG, i.e., the £_10% false negatives that have been reported in several studies (Iritani & Inzana 1988; Andrews et al. 1987; Rippey et al. 1987; Perez et al. 1986; Robinson 1984; Feng & Hartman 1982; Kilian & Bulow 1976).

Since a data base has not been established (or recognized) to clearly determine the reliability of MUG , and since some investigators doubt its effectiveness or reliability, an interim interpretation based on a (MUG+,FC+/MUG-,FC+) presence/absence ratio could give a

general indication of sanitary quality. A low (+/-) MUG ratio would indicate relatively lower levels of E_. col i, as opposed to other non-E^. col i fecal coliforms, and might ease strict interpretation of the fecal coliform test which has caused the declaring of shellfish as unfit for humanconsumption. This information could work together with the current guideline concerning shellfish taken from "approved" waters which are subsequently found to have fecal coliform levels in excess of the regulations. In these cases, ifproper sanitary conditions can be shown to have been in force, excess fecal coliforms are simply looked at as signs of deterioration and not as a direct health threat (NSSP 1988).

Once a good data base for MUG is established,recognized, and its ability to detect E_- coli (EC) confirmed, then the regulations could be revised to reflect coliinstead of fecal coliforms. Should there be concern about a certain percentage of E_. coli not Deing detected on a regular basis, a correction factor could be used and incorporated into an E. coli MPN table. This would be differentiated from the standard fecal coliform MPN table. If, for example, it is found that MUG routinely does not identify 596 of the E. coli, then the MPN values could be altered to take this into account. This would give a margin of safety to the values produced by the FC/MUG test. If this procedure is accepted, then the value of 230 EC/100 gm of sample could be considered

the legal standards for determining the sanitary quality of shellfish and other seafoods. This recommendation has been recently made at the 7*” Annual Meeting of the Interstate Shellfish Sanitation Conference, held at Stamford, CT (July 9-14, 1989). Also, the use of MUG has been recognized for use in testing water samples by the Environmental Protection Agency, starting at the beginning of August 1989 as reported in the Federal Register (June 29,1989).

If the methods are not changed or alternatives mandated, such as relaying or depuration, then there is a possibility that harvesting of shellfish would have to be completely suspended, at least during the warm summer months along the Gulf Coast.

CONCLUSIONS AND RECOMMENDATIONS

This study was predicated on the hypothesis that the Klebsiella sp. being isolated from summer harvested Louisiana oysters could be differentiated from clinical isolates of K. pneumoniae. Based on the DNA-DNA homology results of this investigation, the oyster isolates were >_65% related to a defined strain of K_. pneumoniae. Therefore, it is clear that the oyster isolates in question were K_. pneumoniae and can not be differentiated from clinical isolates of K. pneumoniae using the procedures tested in this investigation, with the exception that many were urease negative. In addition, some of the oyster isolates lost their ability to produce a positive fecal coliform reaction after several passages on BHI agar at 20° to 25°C.

The thermal denaturation and renaturation procedures used in this study are effective but their use as clinical or screening tools would probably be of relatively little use due to the time required. It requires approximately 10 days for DNA results to be generated, from initial isolation of the organisms to the extraction and analysis of the chromosomal DNA. The DNA-DNA homology (renaturation)procedure would, however, be of value should species relatedness information be required.

It is recommended that the standard fecal coliform test be modified to allow it to differentiate E. coli from other

72

non-E^- col i fecal coliforms, in particular 1C. pneumoniae. This can be done by incorporating iOO ug of MUG in EC medium to prevent false positives due to the natural B-glucuronidase present in shellfish tissue . Thereby, the standard should be altered to reflect 230 E_. col i /100 gm instead of fecal coliforms. The test (Table 12) should also be modified by monitoring LST for 24 hours, instead of the current 48 hours. Furthermore, the temperature at which the EC medium is incubated should be increased from 44.5°C to 45.5°C to help reduce the incidence of false positives. If concerns about the reliability of MUG persists, a (+/-) MUG ratio could be implemented to give a general indication of sanitary problems which might exist due to the presence of fecal coliforms other than E. coli.

74

Table 12: Recommended Modified Fecal Coliform Test.

1) Sample preparation using APHA procedures.2) Inoculation of sample into standard LST broth.3) 24 hour incubation of LST at 35°C.4) Inoculation of positive LST tubes into EC medium

containing 10O ug/ml MUG. Second 24 hour incubation of negative LST tubes eliminated.

5) 24 hour incubation of EC at 45.5°C.6) Interpretation of results, fecal coliform and MUG

react ions.7) Report results for appropriate action.

REFERENCES

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VITA

Thomas Elbert Graham was born May 25, 1956 in New Orleans, LA. He graduated from Archbishop Rummel High School in Metairie, LA in May, 1974. He attended Tulane University in New Orleans, LA, majoring in biology and received a Bachelor of Science degree, with honors in biology, in May 1978. In January, 1980 he started working towards a graduate degree in fisheries biology with a minor in microbiology. In January, 1981 he became a full-time research associate in the Department of Forestry and Wildlife Management and continued working part-time towards his degree. He received a Master of Science degree in May, 1983. In May, 1984 he began work as a full time research associate in the Department of Food Science and began degree work in August of that year. He worked part-time towards his degree until February, 1986 at which time he became a full-time student. He is presently a candidate for the degree of Doctor of Philosophy in Food Science.

87

DOCTORAL EXAMINATION AND DISSERTATION REPORT

Candidate: Thomas E. Graham

Major Field: Food S c ie n c e

Title of Dissertation: C om parison o f C l i n i c a l I s o l a t e s o f K l e b s i e l l a pneum oniae toK l e b s i e l l a s p . I s o l a t e d from Summer H a rv e s te d E a s te r n O y s te r s , C r a s s o s t r e a v i r g i n i c a

Approved:

Major Professor and Chairman

•1. J'jk-Dean of the Graduate

EXAMINING COMMITTEE:

RoIkJL toi .

ft*

Date of Examination:

O c to b e r 3 0 , 1989